The three-dimensional structures of functional nucleic acids (RNA/DNA) are the key to understanding their functions, designing targeted drugs, and developing synthetic-biology components. However, the low efficiency of nucleic acid crystallization and the long cycle required for phase determination remain the central bottlenecks constraining this field. We are committed to transforming empirical knowledge into standardized methods and further developing these methods into automated tools, thereby advancing nucleic acid crystallography into a new stage.

Database Development

• Based on the three established data platforms Ribocentre¹ (ribozymes), Ribocentre-switch² (riboswitches), and Ribocentre-aptamer³ (aptamers), we are integrating 50 years of worldwide RNA crystallography data to build the first RNA crystallization methodology database (unpublished).

Two Original Technologies

• GU base-pair engineering: By introducing or removing GU base pairs, we have successfully determined 10 RNA crystal structures^4,5,6.

• The counterintuitive “extension of nonfunctional stems” paradigm: Statistical analysis shows that nonfunctional stem regions in nucleic acids are typically 4–8 base pairs (bp) long. Extending them to 9–12 bp can both increase the probability of crystallization and intrinsically provide a molecular-replacement search model. This strategy has successfully enabled the determination of multiple new functional nucleic acid structures (unpublished).

Finally, I will also discuss which interesting structures we have determined using these efficient methods, what new insights these structures have generated, and what scientific questions they have helped address^7,8,9,10.

Keywords: crystallography; nucleic acids; ribozymes; riboswitches; aptamers

References

[1] Deng, J… Huang, L. Nucleic Acids Research. 2023, 51, D262–D268.

[2] Bu, F.; … Huang, L. Nucleic Acids Research 2024, 52, D265–D272.

[3] Lu, Z.; … Huang, L.; Miao, Z. Nucleic Acids Research 2026, 54, D264–D272.

[4] Ren, Y.; … Huang, L. Nucleic Acids Research 2025, 53 (3), gkae1218.

[5] He, Y.; … Huang, L. Cell Discovery 2025, 10 (1), 128.

[6] Chen, K.; … Huang, L. Nucleic Acids Research 2025, 53 (14), gkaf702.

[7] Deng, J… Huang, L. Nature Chemical Biology 2022, 18 (5), 556–564.

[8] He, Y.; … Huang, L. Nucleic Acids Research 2026, 54 (6), gkag211.

[9] Lin, X.; … Huang, L. Angewandte Chemie 2025, 137 (22).

[10] Wang, J.; … Huang, L.; Tan, J. Chem 2026, 12 (5), 102883.

In the “Dynamics of protein synthesis & RNA decay” lab we are interested in how cells tune protein production by adjusting the stability of messenger RNAs (mRNAs). Each cell in our body expresses around 12,000 different mRNAs at any given time. To control this complex transcriptome—and thus protein synthesis—cells adjust half-lives for individual mRNAs from minutes to several days. Traditionally, the selective degradation of mRNAs has been attributed to the recognition of nucleotide sequence elements by proteins or small RNAs that subsequently recruit decay factors. In our lab, we investigate a distinct, newly emerging paradigm of gene regulation termed “peptide-mediated mRNA decay” (PMD). In PMD, not the mRNA sequence but rather the nascent protein is recognized to trigger degradation of the encoding mRNA during its translation by the ribosome.

This process is exemplified by tubulin mRNAs: Microtubules, built from heterodimers of alpha- and beta-tubulins, control cell shape, mediate intracellular transport and power cell division. The concentration of tubulins is tightly controlled through a post-transcriptional mechanism involving selective and regulated degradation of tubulin-encoding mRNAs. We and others recently demonstrated that degradation is initiated by TTC5, which recognizes tubulin-synthesizing ribosomes and recruits the downstream effectors SCAPER and CCR4-NOT to trigger mRNA decay (Lin et al., 2020, Science; Höpfler et al., 2023, Mol Cell). Subsequently, we identified the first regulatory step of this cascade that controls TTC5 activity (Batiuk, Höpfler, et al., 2024, Nat Commun). Together, our data provide a conceptual framework for peptide-mediated mRNA decay. Building on this, future work will investigate how PMD globally shapes the human transcriptome.

Human RNase 2, also named the eosinophil-derived neurotoxin (EDN), is a secretory protein of the vertebrate-specific RNase A superfamily involved in antiviral and proinflammatory cell response. Identifying ligand-binding pockets in EDN is thus relevant to structure-based drug design. By X-ray crystallography we first identified a conserved site at the protein surface binding to carboxylic anion molecules: malonate, tartrate and citrate. Searching for potential biomolecules rich in anion groups and considering previous report of EDN interaction to glycosaminoglycans, we explored the protein binding to saccharides. EDN crystals were soaked with mono- and disaccharides, and the 3D structures of ten complexes were solved by X-ray crystallography at atomic resolution. We identified protein binding sites to glucose, fucose, mannose, galactose, trehalose, sucrose, N-acetyl-D-glucosamine, N-acetylmuramic acid, and the sialic acid N-acetylneuraminic acid. A main site for glucose, fucose, and galactose was located adjacent to the spotted carboxylic anion site. Secondarily, N-acetylneuraminic acid and mannose shared another close by protein surface region. Overall, we located 17 saccharide ligands that clustered into seven defined sites, outlining a conserved recognition pattern, which was further analysed by molecular modelling. Interestingly, within the RNase A superfamily, we find amphibian RNases that were initially isolated as carbohydrate binding proteins and named as leczymes, combining enzymatic and lectin properties. The present data is the first structural characterization of a mammalian sugar-binding RNase within the family. The results highlight unique EDN residues that mediate sugar specific interactions, of particular interest for a better understanding of the protein physiological role.