Capturing Conformational Dynamics and Transient Interactions

There are four research areas in the Department of Pharmaceutical Chemistry. Capturing conformational dynamics and transient interactions is a research challenge within physical biology.

The challenge

Only a relatively small subset of protein molecules have had their structures experimentally determined (just over 100,000 in the Protein Data Bank as of May, 2014). In the vast majority of cases, this has been done using x-ray crystallography, which requires purifying proteins—with or without bound ligands—coaxing them to form crystals, then shining x-rays through them and analyzing how they alter (diffract) the beams.

But that method traditionally captures static snapshots, whereas proteins are flexible molecules that, beyond their basic chemical configurations, take on different shapes (conformations), including those essential to carrying out their functions.

It is this shape-shifting capability that allows proteins to be regulated. Knowing which conformations predominate and their rates of interchange is vital to understanding biological processes and potentially designing therapeutic interventions. Indeed, some proteins, including those dysregulated in cancers, exist in both active and inactive conformations that have distinct affinities for small molecules and thus major implications for drug design.

Also, some proteins are too flexible to be crystallized in their full-length form (e.g., infectious prions that cause neurological diseases) or have sub-structures that are altered by crystallization from the shapes they would take in vivo (e.g., loops that connect secondary structures and which often contain catalytic or active sites that are typical targets of therapeutic agents).

Finally, many important binding interactions between proteins and/or with ligands are weak and thus brief (to quickly meet biological needs), making them difficult to capture via crystallization. Such interactions, as well as shifts in conformation, can occur on times scales of thousands to millionths of seconds, yet crucially alter their activity.

Examples of our methods and research include

Department scientists apply a variety of biophysical methods to detect and analyze the structures, conformational dynamics and interactions of macromolecules such as proteins and nucleic acids (e.g., RNA) as well as small molecules and metabolites. These include:

  • X-ray crystallography—to determine atomic-level molecular architecture, including specific active (binding) and/or inactive conformations and the binding of catalytically essential metal ions.
  • Small-angle X-ray scattering (SAXS)—x-ray scattering in solution that generates coarser measures of molecular envelopes rather than structural details, thus detecting major changes in conformation.
  • Fluorescence resonance energy transfer (FRET)—measures of energy transfer efficiency between two fluorescent labeling compounds (fluorophores) in order to determine distances between them, providing measures of structure and conformational change.
  • Nuclear magnetic resonance (NMR) spectroscopy—which sensitively detects flexible macromolecules and structures (e.g., transmembrane proteins, RNA, multi-domain proteins) in solution rather than in a crystallized form, thus capturing them as they cycle through ensembles of conformations (dynamics), in transient interactions with other molecules, and as in vitro confirmation of binding activity and locations in a non-crystallized state.

In addition to providing information on individual proteins and their substructures (domains, active sites), their conformational dynamics, and specific sites of weak/transient interactions, these biophysical methods measure the results of experiments that apply the modern tools of molecular and chemical biology, such as site-directed mutagenesis, enzyme inhibition, and kinetic analysis (e.g., showing how changes in specific residues alter the rate of enzyme catalytic action or determining binding mode of a drug to an allosteric site).

Notably, the department is home to the state-of-the-art UCSF NMR Laboratory, used by department, School, UCSF and qualified outside researchers.

NMR laboratory

Department-based UCSF Nuclear Magnetic Resonance (NMR) laboratory

NMR spectroscopy resolves molecular function by detecting specific atomic nuclei, such as those of ubiquitous hydrogen atoms, which spin like tops with distinctive wobbles (precessions) when placed in a magnetic field. (NMR spectrometers generate such fields using powerful super-cooled, super conducting magnets.)

Each atom’s rate of precession (frequency) can also be altered not just by the overall magnetic field generated by the spectrometer, but additionally by local fields from the spinning nuclei in adjacent atoms and molecular structures.

NMR spectrum

NMR spectrum of hexaborane (B6H10) molecule showing peaks shifted in frequency, which give clues to the molecular structure.

When they are subjected to short bursts (pulses) of radio waves, precession frequencies can be detected, amplified, and represented in charts called spectra. A spectrum gives a list of precession frequencies, reporting on the magnetic environment of a specific hydrogen. This will change if a protein changes its conformation or is bound by another molecule.

Thus NMR is a uniquely sensitive way to study both flexible molecular structures and on-going shifts in a conformation (dynamics). The method can detect weak/transient interactions between molecules, since there will be a change in the precession frequency at their binding site with changes in the magnetic environment as the two chemical groups interact. NMR here can detect such dynamics and interactions occurring as quickly and briefly as mere billionths of seconds (nanoseconds).

Analyzing enzyme dynamics and interactions during a key regulatory event in cell biology

The central dogma of molecular biology states that DNA makes RNA, which makes proteins, and proteins carry out the functions of living organisms. Specifically, messenger RNA (mRNA) carries the blueprints for making specific proteins from encoding genes in the cell nucleus for processing by ribosome factories in the cytoplasm.

Such an essential process of life is by necessity highly regulated. For example, there’s a quality control process disassembling mRNAs (via enzymatic degradation, also known as decay) that are truncated or malformed and would thus generate mutant proteins (nonsense-mediated decay). Also, certain mRNAs are degraded as the end results of sequential protein interactions (pathways) in order to maintain cellular homeostasis, to limit production of a particular protein, or so the cell can make more of other proteins to respond to needs in an animal’s development, cell differentiation/proliferation, and stress/immune responses.

Gene expression can thus be controlled at the level of mRNA (transcript) stability—and failures in such regulation may yield cancers and other diseases.

Department scientists use a variety of biophysical methods, including NMR, to guide and detect the results of biochemical and genetic mutagenesis experiments that dissect the penultimate, irreversible step in numerous eukaryotic mRNA decay pathways. In that step, the mRNA decapping enzyme 2 (Dcp2) removes a structural cap (7-methylguanosine or m7G) via hydrolysis at one end (five prime terminal or 5′ end) of an mRNA, exposing a 5′-monophosphate that is recognized by 5′-to-3′ exonucleases—enzymes that cleave the mRNA’s nucleotides apart starting from that end.

Using yeast as model organisms (key protein domains are conserved in humans) researchers here are discovering the details of how this mRNA decapping is carried out. In particular, they explore how the chemical reaction rate (enzyme kinetics) of this key step in mRNA decay is affected and controlled by rapid changes in the shape of Dcp2 (conformational dynamics) that bring two domains together to form the enzyme’s active site and the weak, very fast, yet critical interactions between Dcp2 and both transcript nucleotides and co-activator proteins in a transient enzyme-substrate complex.

process of mRNA decapping by enzyme

Model of decapping catalysis by Dcp2, which proposes that the above conformational change is used as a point of control by decapping activators.
(a) Dcp2 exists in a conformational equilibrium between open and closed forms.
(b) RNA (with m7G magenta-hexagonal cap) binds to the open form.
(c) After binding the enzyme closes over the RNA substrate, a step that may be enhanced by co-activators.
(d) Following closure, the RNA cap is removed by hydrolysis.
(e) The RNA is then released and subject to further degradation (exonucleolysis) by 5'-3' nucleases.

Experimental findings by department scientists and their collaborators using NMR spectroscopy and other methods include

open and closed forms of Dcp2

Open and closed forms of Dcp2 with co-activator Dcp1 binding the regulatory domain

Characterizing key decapping enzyme conformational cycling

Revealing that yeast Dcp2 in solution exists in a conformational equilibrium. It has a dumbbell-shaped (bilobed) N-terminus—one lobe is the catalytic domain shared by the Nudix enzyme superfamily that hydrolyzes nucleotide diphosphates like those in the mRNA cap and the other lobe is a regulatory site (which binds the essential coactivator Dcp1). The enzyme is thought to cycle between an “open” (mostly inactive) and closed (active) forms, in which these two structural domains come together in an active site to degrade mRNA far more efficiently. Indeed, in the absence of ligand, department-based NMR spectroscopy finds Dcp2 cycles from open to closed to open states roughly every 500 microseconds or approximately 2000 times a second.

Quantifying effects of conformational shifts on reaction rates

Determining that the rate at which Dcp2 catalyzes mRNA decapping is dramatically increased when the enzyme is in a “closed” as opposed to an “open” state—when the catalytic and regulatory domains come together upon cap recognition (after substrate binding) to form a composite active site. This site sandwiches the transcript cap, which department NMR spectroscopy shows binds with conserved residues on both domains. Findings by department scientists also indicate the regulatory domain helps position the cap for catalysis by the opposing Nudix domain. When researchers here deleted the regulatory domain’s cap-binding site, catalysis was reduced about 100-fold and the closed conformation is blocked.

Measuring the regulatory effects of co-activator binding

Showing that binding of Dcp2’s regulatory domain by the coactivator Dcp 1 increases catalysis by a factor of ten in itself and by the latter binding coactivators such as Edc1 or Edc2 increases decapping 1,000-fold. This suggests a regulatory model in which interactions with protein cofactors stabilize the Dcp2’s mutable composite active site to greatly increase the rate of decapping.


A model of mRNA recognition and decapping: The complex of Dcp2 (green) and Dcp1 (yellow) in the background is bound to the RNA substrate body (purple strand) yet remains in an open conformation. But the Dcp1/2 complex in the foreground, with access to the 5’ end cap structure (red and purple balls and sticks) assumes a closed and greatly activated conformation.

Determining how decapping enzyme recognizes its mRNA substrate

Clarifying the molecular mechanisms of mRNA recognition by the Dcp2 decapping complex. Revealing (via NMR spectroscopy and kinetic analyses) that activation of the complex (including the recruitment of Dcp2 coactivators) is affected by access to nucleotides in the RNA body next to the 5’ end. NMR was required because Dcp2 interaction with the cap is weak but specific and requires RNA-binding to a dynamic interface—a channel intersecting the catalytic and regulatory domains. It is thought that this restricts decapping to transcripts with exposed 5’ends as a way of preventing their premature degradation.

Linking active site structure and conformational dynamics to catalysis

Analyzing how Dcp2 active site conformational dynamics, enhanced by the binding of Dcp1 and other coactivators, affect catalytic chemistry. Department scientists used x-ray crystallography of the catalytic domain to demonstrate the binding of an essential metal ion by three specific glutamic acid residues. They also employed pH-dependent enzyme kinetics experiments to identify several key active site residues involved in the acid/base catalysis of decapping. In addition, NMR and computer modeling was applied to determine that a conserved metal binding loop on the catalytic domain (location of a conserved glutamic acid residue crucial for decapping activity) shifts conformation during catalysis.

Revealing key residue for conformational transition and coactivation

Using NMR spectroscopy of enzyme kinetics, department researchers showed that a specific conserved Dcp2 tryptophan residue acts as a vital gatekeeper of the open-to-closed conformational transition, promoting formation of the composite active site and enabling Dcp1 and its coactivators to further enhance catalysis. Indeed, experimental mutation of the residue, Trp43, eliminates Dcp2 conformational dynamics, blocks regulatory domain cap binding, and even reduces the enhancement of enzymatic activity by Edc1, which binds to Dcp1 of the decapping complex and increases the complex’s catalytic efficiency up to 3,000 times. This work further supports the model in which such cofactors regulate mRNA decapping via stabilization of Dcp2’s composite active site.


NMR spectra reveal large, global conformational differences between wild-type yeast Dcp2 (magenta)—specifically the N-terminal residues encompassing the regulatory and catalytic domains that form the composite active site—and a Dcp2 (black) in which a conserved gatekeeper amino acid, tryptophan43, is mutated to alanine. The wild-type Dcp2 samples multiple conformations in solution, while the mutant quenches dynamics vital to catalysis and locks the protein into one conformation.

Characterizing locations, effects of additional decapping co-activators

Using NMR spectroscopy and mutational studies, researchers here analyzed the locations and kinetic effects of additional decapping activators. Along with collaborators, they showed that the enhancers of decapping protein family (Edc1 and -2) in S. cerevisiae yeast bind via a proline-rich sequence to a site on Dcp1 and stimulate decapping 1000-fold. Further Edc1 increases the catalytic efficiency of the Dcp1-Dcp2 complex both by enhancing the catalytic step and increasing mRNA binding. Other labs in the field have built on this to show the same type of interactions are used in mRNA quality control pathways in humans during nonsense-mediated decay (NMD).