About

Sarah Woodson is the T.C. Jenkins Professor of Biophysics at Johns Hopkins University. She received her PhD in Biophysical Chemistry from Yale University in 1987, where she studied under Donald Crothers, and completed postdoctoral research in the laboratory of Thomas Cech at the University of Colorado Boulder from 1987 to 1990. Her research group focuses on studying how RNA molecules fold into specific three-dimensional structures and how RNA and proteins within cellular complexes such as the ribosome come together. Her work employs biophysical methods including X-ray scattering, single-molecule fluorescence, and X-ray hydroxyl radical footprinting, which she has pioneered for RNA analysis. Dr. Woodson's research explores the physical processes of RNA folding, the assembly of ribosomes, and the role of small non-coding RNAs and the chaperone protein Hfq in genetic regulation. Her studies on RNA folding pathways, the stability of RNA structures, and the mechanisms of ribosome assembly contribute to understanding cellular function, disease mechanisms, and evolutionary processes. She has received numerous awards, including the Pew Scholar Award in Biomedical Sciences and the Camille Dreyfus Teacher-Scholar Award, and was elected an AAAS Fellow in 2010. She also served as President of the RNA Society from 2016 to 2017.

Research topics

• Biology
• Genetics
• Computational biology
• Cell biology
• Physics
• Microbiology

Selected publications

• Elucidating structure–function relationships in the mammalian nucleolus

  Nature Reviews Molecular Cell Biology · 2026-05-19

  article
  PublisherDOI

• Metastable structure of Bacillus subtilis glmS ribozyme permits turnover by RNase J1

  SSRN Electronic Journal · 2025-01-01

  preprintOpen accessSenior author
  PublisherDOI

• Competition for Hfq drives kinetic selection of mRNA targets by <i>Escherichia coli</i> small noncoding RNAs

  Proceedings of the National Academy of Sciences · 2025-07-10 · 2 citations

  articleOpen accessSenior authorCorresponding

  bacteria in response to diverse stimuli in the environment and the host. Most sRNAs regulate mRNA expression by directly base pairing with complementary sites in the target mRNA with the help of the chaperone protein Hfq. sRNAs and Hfq must rapidly search hundreds of candidate mRNAs for matched (cognate) targets while discriminating against noncognate targets. Here, we use single-molecule fluorescence microscopy to directly observe how cognate and noncognate mRNAs bind immobilized sRNA-Hfq. The results show that initially unstable sRNA-Hfq-mRNA complexes either dissociate within seconds by ejecting one of the two RNAs, depending on their interactions with Hfq, or are stabilized by sRNA-mRNA base pairing. Cognate mRNAs are more likely to form long-lived sRNA-Hfq-mRNA complexes, even in the presence of competing RNA. Active competition for the mutual Hfq chaperone introduces a kinetic barrier to RNA colocalization that is resolved by base pairing, driving the accumulation of cognate sRNA-mRNA interactions while eliminating noncognate interactions.

  PublisherDOI

• Disassembly of unstable RNA structures by an <i>E. coli</i> DEAD-box chaperone accelerates ribosome assembly

  Nucleic Acids Research · 2025-02-08 · 5 citations

  articleOpen accessSenior author

  Ribosome synthesis in bacteria is coupled with transcription of the pre-ribosomal RNA (pre-rRNA), which must fold and assemble with 20 or more ribosomal proteins. In vitro, the Escherichia coli pre-16S rRNA misfolds during transcription, delaying stable binding of ribosomal protein uS4 that nucleates assembly of the 16S 5' domain. Using single-molecule fluorescence microscopy, we show that the DEAD-box protein CsdA (DeaD) strongly accelerates uS4 binding by facilitating proper folding of the nascent rRNA. Unstable RNA structures are unfolded by CsdA, whereas stable RNA structures resist unwinding. We show that CsdA unfolding becomes less frequent as more ribosomal proteins add to the complex. The results demonstrate that disassembly of unstable, nascent RNA-protein complexes by chaperones fuels the search for native structure. We propose that general chaperones create a gradient of disassembly that steepens the hierarchy of proper protein addition until late assembly intermediates escape unwinding and commit to 30S maturation.

  PublisherDOI

• Stick-slip unfolding favors self-association of expanded HTT mRNA

  Nature Communications · 2024-10-09 · 2 citations

  articleOpen accessSenior author

  In Huntington's Disease (HD) and related disorders, expansion of CAG trinucleotide repeats produces a toxic gain of function in affected neurons. Expanded huntingtin (expHTT) mRNA forms aggregates that sequester essential RNA binding proteins, dysregulating mRNA processing and translation. The physical basis of RNA aggregation has been difficult to disentangle owing to the heterogeneous structure of the CAG repeats. Here, we probe the folding and unfolding pathways of expHTT mRNA using single-molecule force spectroscopy. Whereas normal HTT mRNAs unfold reversibly and cooperatively, expHTT mRNAs with 20 or 40 CAG repeats slip and unravel non-cooperatively at low tension. Slippage of CAG base pairs is punctuated by concerted rearrangement of adjacent CCG trinucleotides, trapping partially folded structures that readily base pair with another RNA strand. We suggest that the conformational entropy of the CAG repeats, combined with stable CCG base pairs, creates a stick-slip behavior that explains the aggregation propensity of expHTT mRNA.

  PublisherOA PDFDOI

• Stick-slip unfolding favors self-association of expanded <i>HTT</i> mRNA

  bioRxiv (Cold Spring Harbor Laboratory) · 2024-06-03

  preprintOpen accessSenior author

  ABSTRACT In Huntington’s Disease (HD) and related disorders, expansion of CAG trinucleotide repeats produces a toxic gain of function in affected neurons. Expanded huntingtin (exp HTT ) mRNA forms aggregates that sequester essential RNA binding proteins, dysregulating mRNA processing and translation. The physical basis of RNA aggregation has been difficult to disentangle owing to the heterogeneous structure of the CAG repeats. Here, we probe the folding and unfolding pathways of exp HTT mRNA using single-molecule force spectroscopy. Whereas normal HTT mRNAs unfold reversibly and cooperatively, exp HTT mRNAs with 20 or 40 CAG repeats slip and unravel non-cooperatively at low tension. Slippage of CAG base pairs is punctuated by concerted rearrangement of adjacent CCG trinucleotides, trapping partially folded structures that readily base pair with another RNA strand. We suggest that the conformational entropy of the CAG repeats, combined with stable CCG base pairs, creates a stick-slip behavior that explains the aggregation propensity of exp HTT mRNA.

  PublisherOA PDFDOI

• A DEAD-box chaperone promotes ribosome assembly by iteratively refolding newly transcribed rRNA

  Biophysical Journal · 2024-02-01

  articleOpen accessSenior author
  PublisherOA PDFDOI

• RNA compaction and iterative scanning for small RNA targets by the Hfq chaperone

  Nature Communications · 2024-03-07 · 14 citations

  articleOpen accessSenior author

  RNA-guided enzymes must quickly search a vast sequence space for their targets. This search is aided by chaperones such as Hfq, a protein that mediates regulation by bacterial small RNAs (sRNAs). How RNA binding proteins enhance this search is little known. Using single-molecule Förster resonance energy transfer, we show that E. coli Hfq performs a one-dimensional scan in which compaction of the target RNA delivers sRNAs to sites distant from the location of Hfq recruitment. We also show that Hfq can transfer an sRNA between different target sites in a single mRNA, favoring the most stable duplex. We propose that compaction and segmental transfer, combined with repeated cycles of base pairing, enable the kinetic selection of optimal sRNA targets. Finally, we show that RNA compaction and sRNA transfer require conserved arginine patches. We suggest that arginine patches are a widespread strategy for enabling the movement of RNA across protein surfaces.

  PublisherOA PDFDOI

• Late consolidation of rRNA structure during co-transcriptional assembly in <i>E. coli</i> by time-resolved DMS footprinting

  bioRxiv (Cold Spring Harbor Laboratory) · 2024-01-10 · 5 citations

  preprintOpen accessSenior authorCorresponding

  Abstract The production of new ribosomes requires proper folding of the rRNA and the addition of more than 50 ribosomal proteins. The structures of some assembly intermediates have been determined by cryo-electron microscopy, yet these structures do not provide information on the folding dynamics of the rRNA. To visualize the changes in rRNA structure during ribosome assembly in E. coli cells, transcripts were pulse-labeled with 4-thiouridine and the structure of newly made rRNA probed at various times by dimethyl sulfate modification and mutational profiling sequencing (4U-DMS-MaPseq). The in-cell DMS modification patterns revealed that many long-range rRNA tertiary interactions and protein binding sites through the 16S and 23S rRNA remain partially unfolded 1.5 min after transcription. By contrast, the active sites were continually shielded from DMS modification, suggesting that these critical regions are guarded by cellular factors throughout assembly. Later, bases near the peptidyl tRNA site exhibited specific rearrangements consistent with the binding and release of assembly factors. Time-dependent structure-probing in cells suggests that many tertiary interactions throughout the new ribosomal subunits remain mobile or unfolded until the late stages of subunit maturation.

  PublisherOA PDFDOI

• Optimized periphery-core interface increases fitness of the <i>Bacillus subtilis glmS</i> ribozyme

  Nucleic Acids Research · 2024-09-25 · 2 citations

  articleOpen accessSenior author

  Like other functional RNAs, ribozymes encode a conserved catalytic center supported by peripheral domains that vary among ribozyme sub-families. To understand how core-periphery interactions contribute to ribozyme fitness, we compared the cleavage kinetics of all single base substitutions at 152 sites across the Bacillus subtilis glmS ribozyme by high-throughput sequencing (k-seq). The in vitro activity map mirrored phylogenetic sequence conservation in glmS ribozymes, indicating that biological fitness reports all biochemically important positions. The k-seq results and folding assays showed that most deleterious mutations lower activity by impairing ribozyme self-assembly. All-atom molecular dynamics simulations of the complete ribozyme revealed how individual mutations in the core or the IL4 peripheral loop introduce a non-native tertiary interface that rewires the catalytic center, eliminating activity. We conclude that the need to avoid non-native helix packing powerfully constrains the evolution of tertiary structure motifs in RNA.

  PublisherOA PDFDOI

Recent grants

• NIH Grant R01GM046686

  NIH · $5.5M · 2016

• Time-Resolved Hydroxyl Radical Footprinting of RNA

  NIH · $6.4M · 1999–2021

• Assembly Mechanisms of RNA-Protein Complexes for Genetic Control

  NIH · $3.9M · 2020–2030

• Assembly Mechanisms of RNA-Protein Complexes for Genetic Control

  NIH · $473k · 2020–2025

• Hfq RNA Chaperone and the Mechanism of RNA-Dependent Regulation

  NIH · $1.3M · 2016–2021

Frequent coauthors

• Robert M. Briber

  University of Maryland, College Park

  152 shared

• D. Thirumalai

  112 shared

• Joon Ho Roh

  103 shared

• Liang Guo

  First Affiliated Hospital of Henan University

  67 shared

• Duncan Kilburn

  Pacific Biosciences (United States)

  64 shared

• Alexei P. Sokolov

  Oak Ridge National Laboratory

  42 shared

• Gokhan Caliskan

  42 shared

• Prashanth Rangan

  Icahn School of Medicine at Mount Sinai

  39 shared

Education

• postdoctoral, Chemistry and Biochemistry

  University of Colorado Boulder

  1990

• PhD, Chemistry

  Yale University

  1987

• B.A.

  Kalamazoo College

  1982

Awards & honors

• Pew Scholar Award in the Biomedical Sciences (1993)
• Camille Dreyfus Teacher-Scholar Award (1995)
• AAAS Fellow (2010)
• President of the RNA Society (2016-2017)