Mar 132009
ResearchBlogging.orgI’ve mentioned urea and guanidinium (Gdm) before on this blog, usually with reference to questions about their mechanism of action. These small molecules cause proteins to denature, or lose their higher levels of structure and become unfolded chains. The complete unfolding of a protein typically requires a fairly high concentration of denaturant, almost always more than 1M, and the explanation for this is that the denaturant molecules preferentially associate with the polypeptide chain with low affinity. In a recent issue of PNAS, a paper from Walter Englander argues that urea, but not guanidinium, associates with the backbone of the protein via hydrogen-bonding interactions.

Lim et al. reached this conclusion using hydrogen-exchange experiments. Amide nitrogens in proteins freely exchange their covalently-bound hydrogens (protons) with the surrounding water. The rate of this process can be measured (among other ways), by placing a protonated amide group into a deuterated solvent and tracking the decline in proton signal by NMR; this is called an HX experiment. In the case of a folded protein chain the observed rate will depend on the intrinsic chemistry of the particular amide and the stability of the protein structure, because this structure excludes water from the backbone and makes hydrogen bonds that lock the protons in place. Rather than deal with all of that, the authors used a small peptide mimic that (probably) has no complex structure. This had the additional advantage that the simple spectrum could be tracked by 1-D NMR, substantially increasing the time-resolution of the measurements. The authors measured the rates as they varied the pH — because we’re talking about D2O, it’s called the pD instead — and added various cosolutes that are known to denature or stabilize protein folds.

As expected, the dialanine itself had a V-shaped rate profile in these HX experiments, with a minimum at a pD of 4. The hydrogen exchange reaction can be catalyzed by acid or base, so the rate increases as you go up or down in pD from this minimum. When urea was added to the solution, the authors found that acid-catalyzed HX accelerated while base-catalyzed HX decelerated. The most reasonable explanation for the latter result is that a hydrogen bond between the carbonyl of urea and the amide proton protects it from water attack. The authors do some mathematical modeling to establish that the effect on rate reflects a bonding association between the peptide and urea, not just random collisions or thermodynamically neutral associations.

The acid-catalyzed result is interesting, because in theory one would expect that urea would accelerate acid-catalyzed HX more than it actually does, because under acidic conditions it can accept a hydrogen from the amide nitrogen. While there are some confounding factors, the most likely explanation for this result is that the NH2 groups of urea form hydrogen bonds to the carbonyl of the peptide. Because acid catalysis of HX hinges on the favorability of protonating this carbonyl, a hydrogen bond would be expected to reduce the HX rate. The authors argue that the ability of urea to serve as an acid catalyst is therefore mitigated by its propensity to bind to the carbonyl.

The formation of hydrogen bonds between urea and the peptide group meshes well with evidence that it denatures proteins through interactions with the backbone, some of which I have mentioned before. From HX experiments under native conditions we know that even a folded protein chain regularly undergoes excursions from its water-excluded, hydrogen-bonded state. Urea may bind to the backbone during these fluctuations, preventing or slowing a return to the folded structure.

Lim et al. also tested a number of other cosolutes, and found that none of them had a similar effect on the HX rate. In the case of the stabilizing molecules (glycerol, sorbitol) this is entirely expected, as their action cannot be explained in terms of a preferential association with the backbone anyway. The surprise concerns guanidinium, which is a more powerful denaturant than urea. The authors noted that Gdm has a small effect on the rate, but not in a pD-dependent way, and one that was little different from an equivalent concentration of NaCl (ordinary table salt). Gdm has no groups that can hydrogen bond to the amide, so the absence of an effect on base-catalyzed HX is expected. However, it should be possible for guanidinium to hydrogen-bond to the carbonyl, so it should seemingly have an effect on acid catalysis. This is not in fact the case.

The authors note that existing evidence does not support the idea that Gdm forms hydrogen bonds with water (although urea is known to do so). Lim et al. suggest instead that the planar Gdm molecule forms favorable stacking interactions with other planar groups. These include the peptide bond and several side chains. They argue that the stacking of Gdm with these groups pries the protein apart without requiring hydrogen bonds.

As a means to investigate diseases that result from protein misfolding, many groups are now trying to structurally characterize the unfolded state of protein molecules. Many of these experiments model the in vivo denatured state by using chemical denaturants such as urea or Gdm. The possibility that direct interactions between the denaturant and the protein will give rise to experimental artifacts should be taken seriously. Urea’s promiscuous formation of hydrogen bonds with the backbone, itself, and water, may give rise to loose networks of hydrogen-bonded molecules that act to condense the chain. By contrast, Gdm’s stacking effect will likely act to artificially extend the chain by steric obstruction. Because of the difference in these mechanisms, it may be of value to cross-validate findings from structural studies on unfolded states by repeating experiments with alternative denaturants.

Lim, W., Rosgen, J., & Englander, S. (2009). Urea, but not guanidinium, destabilizes proteins by forming hydrogen bonds to the peptide group Proceedings of the National Academy of Sciences, 106 (8), 2595-2600 DOI: 10.1073/pnas.0812588106

Feb 272009
ResearchBlogging.orgDecades of studies involving extensive mutagenesis of proteins and protein domains have impressed on us the idea that the folded tertiary structures of proteins are fairly resilient. While a particular mutation may abolish function by directly ablating a key chemical group, it is rare for a single mutation, or even a group of several mutations, to significantly change the overall conformation of a folded polypeptide chain. When a major change does result, it often takes the form of complete denaturation. Because of this, it may seem that protein folds occupy stable islands in sequence space, surrounded by a sea of sequences that form random coils or molten globules. However, there is some evidence that this view is mistaken, that substantially divergent structures may have very similar sequences. A paper recently published in PNAS adds to this view by describing a major change in the structure of a PAS domain resulting from 1-3 mutations.

PAS is a large family of protein-protein interaction domains contained in many signaling proteins. Although many of them have cofactors that modulate their binding, some PAS domains are constitutively active, which is the case for the domain under study here, the PAS-B domain from one of the founding members of the family, the aryl hydrocarbon receptor nuclear transporter (ARNT). The PAS-B domain was believed to dimerize with PAS domains from other proteins through one side of its β-sheet, and as a consequence Evans et al. decided to make several mutations on the outer surface of the sheet and monitor their effects on dimer formation.

One of these mutations, Y456T, had a strange effect on the domain’s NMR spectrum: about 30 new peaks showed up in the 1H-15N HSQC. Because this spectrum should show a single peak for each chemically unique proton-nitrogen pair, this result suggests that the pure proteins in the magnet exist in two distinct conformations. By lowering the temperature and repurifying the protein, Evans et al. were able to mostly separate these two conformations from each other. However, over time these conformationally purified samples became heterogeneous again, which means that the conformations can freely interconvert. This process was very slow, however — too slow to be detected using NMR relaxation techniques. From monitoring the HSQCs the authors concluded that the time constant for interconversion in the mutant is on the order of 16 hours.

Intrigued by what they were seeing, Evans et al. made additional mutations to ARNT PAS-B and found that the proportion of protein in each structure can be adjusted within a wide range by mutation. Using a triple-mutant system they were able to drive about 99% of the proteins into the new conformation. Using this mutant the authors were able to assign the resonances in the HSQC spectrum and solve the structure using NOEs. They learned that the chemical shift changes in the mutant are quite widespread, as you can see in the figure I have shamelessly stolen (left) for your benefit. In this figure the chemical shift changes are mapped onto their structure of the new conformation using color, ranging from green (very little or no change) to red (significant changes) — the sites of the three mutations are shown. As you can see, the chemical shift effect is widespread, covering almost the whole β-sheet of the protein and reaching to the helix on the opposite side.

Closer analysis of the structure shows why this is so. The strand of the β-sheet on which Y456 is situated has shifted its register by 3 spots. This has two effects. The first is that all of the hydrogen bonds involving that strand must be broken, at a substantial energetic cost. This is probably the reason the interconversion process is so slow. Moreover, because the register shifts by an odd number of residues, the strand must flip over, exposing to solvent the residues that were buried in the original structure, while burying the residues that were previously solvent-exposed. Although the backbone and side chain orientations in the strand overlay reasonably well between the two structures, the chemical groups available for interactions are completely different. Unsurprisingly, this alternate structure has very low affinity for its natural targets — titration experiments showed that the alternate conformation bound to its partner from hypoxia inducible factor at least 100x worse than the native structure.

Of course, in living cells with a wide variety of surfaces to bind to, the mutant PAS-B might find an alternative partner for which it has high affinity. Studies that attempt to understand protein-protein interfaces from an engineering or evolutionary perspective typically adopt the assumption that point mutations have little effect beyond adding a particular functional group here or there. This study indicates that this attitude underestimates the ability of point mutations to radically remodel interface surfaces. While a Y→T mutation may not seem particularly conservative, a Y→S mutation has similar effects and requires only a single nucleotide base change. It is not inconceivable that this alternate conformation could be reached in vivo, potentially giving rise to completely novel protein-protein interactions.

One might well wonder how common rearrangements of this kind are likely to be. As the authors point out, structural plasticity in the β-sheet is likely to be a common feature of PAS domains, making it difficult to assess whether this kind of mutational effect is widespread in other folds. However, the authors cite several examples of β-strand register shifts in other proteins. In addition, our decades of alanine-scanning mutagenesis have little to tell us about how common these kinds of rearrangements are, for two main reasons. First, the structural effects strongly depend on what a given residue is mutated to (see Table 1); had Evans et al. been content to leave things at alanine mutations they would never have detected this effect. Second, widely-applicable techniques for sensitively detecting small populations of alternate conformations have not been available until recently.

While the conformational transition induced by the Y456T mutation preserves the protein’s overall fold and stability, it rearranges the hydrogen bonding contacts of the main β-sheet and shortens a loop. Obviously this is not as dramatic a change as found in lymphotactin. However, this alternate structure has significant consequences for PAS-B function. Unexpectedly, this single point mutation can radically remodel the PAS-B binding surface. Moreover, this result adds to the evidence that new structures (even in the context of known folds) may be accessed with only a few changes in amino acid sequence, without any need to detour through molten-globule intermediates.

M. R. Evans, P. B. Card, K. H. Gardner (2009). ARNT PAS-B has a fragile native state structure with an alternative β-sheet register nearby in sequence space Proceedings of the National Academy of Sciences, 106 (8), 2617-2622 DOI: 10.1073/pnas.0808270106

 Posted by at 2:00 AM
Oct 172008
ResearchBlogging.orgProtein stability studies that rely on the use of cosolutes to effect chemical denaturation typically use either urea or the guanidinium ion (Gdm+). Both of these chemicals unfold proteins through a process involving multiple, low-affinity binding events, but guanidinium has the larger effect. While this could be attributed simply to the strength of the interactions, it could also be a result of other chemical properties. In an upcoming paper in the Journal of Physical Chemistry, a team of researchers from the University of Pennsylvania suggest that a reorganization of hydrogen bonds in water may be partially responsible for its greater denaturing power.

The reason water is a liquid rather than a gas is the existence of significant networks of transient hydrogen bonds between water molecules. Typically, water forms these bonds proficiently at a wide range of angles, which means an energetic benefit without a high entropic cost. The range of bond angles in use can be tested by assessing their vibrational frequency using infrared (IR) spectroscopy. The core of this paper comes from an experiment Scott et al. perform in which they measure the dependence of these vibrational frequencies on the concentration of Gdm+ and the temperature of the solution.

They find that the presence of Gdm+ significantly alters the IR spectrum of water at high temperatures, shifting the overall peak to a lower wavenumber and causing the appearance of a shoulder around 3300 cm-1, near the main peak of the ice spectrum (Figure 2). The effect of the Gdm+ ion appears to be less at lower temperatures; as a result the appearance of the water spectra changes less at high Gdm+ concentrations than in the absence of the solute. These spectral characteristics are not observed when another positive ion (potassium) is used instead of Gdm+.

The authors interpret these changes in the IR spectrum as an increase in the number of short, linear hydrogen bonds, and back this up with quantum chemical simulations. If this is correct, then it implies that Gdm+ rearranges the hydrogen bonding network of water, changing the structure to emphasize strong hydrogen bonds of a particular geometry. Moreover, this change in structure appears to be relatively resistant to changes in temperature. This is significant because the creation of protein tertiary structure is thought to be a mechanism by which systems avoid forming highly-structured networks of water hydrogen bonds. If Gdm+ induces significant water structuring, might that entropically favor the unfolding of proteins?

Because urea does not appear to induce this kind of change in water structure, it should be possible to test this experimentally. The effect of Gdm+ is clearly most pronounced at higher temperatures. So one could perform a comparison of the temperature dependence of Gdm+ and urea denaturation of a protein. Proteins tend to be more stable at lower temperatures, as a result we would expect the denaturant concentration at which half of the protein population is unfolded (D1/2) to increase as the temperature decreases. If the structuring of water is significant, then the change in D1/2 should be larger for Gdm+ than for urea, because Gdm+ is decreasing in effectiveness as the temperature goes down. The direct interaction may also be temperature-dependent, but it may be possible to control for this by measuring the energy released when guanidine interacts with an intrinsically unfolded protein.

Unfolding experiments typically are not interpreted in a way that that depends on the precise mechanism by which Gdm+ breaks down protein structure. Nonetheless, the specifics of this process are important if we want to use chemically denatured states as models of in vivo unfolded states. As we gain a better understanding of in vivo water structure, the characteristics of denaturant solvation may be an important consideration in experimental design.

J. Nathan Scott, Nathaniel V. Nucci, Jane M. Vanderkooi (2008). Changes in Water Structure Induced by the Guanidinium Cation and Implications for Protein Denaturation Journal of Physical Chemistry A DOI: 10.1021/jp8058239

 Posted by at 1:00 AM
Mar 262008
ResearchBlogging.orgWhile reports of my man-crush on Brian Volkman are in general much exaggerated, it is true that I adore one of the systems he studies, the bizarre chemokine lymphotactin. In case you couldn’t guess from past posts here, I am endlessly fascinated by this protein, and so I was very happy to finally see his latest paper on the subject in today’s feed-dump from PNAS. Previously published research out of Brian’s group indicated that lymphotactin adopted two totally different structures under different solution conditions. The new paper provides high-resolution structures of the non-chemokine fold and demonstrates that the structures have distinct activities, both of which are essential for full lymphotactin function in vivo.

Lymphotactin is a chemokine, a small protein which has the property of binding to molecules in the extracellular matrix (ECM) such as long polysaccharides, and also of activating certain G-protein coupled receptors (GPCRs). A previously-solved structure of lymphotactin (left) displayed a normal chemokine fold (explore this structure at the PDB), which is designated Ltn10. However, in order to get this structure by NMR, the Volkman lab had to either engineer in a second disulfide bond (as in this structure), or acquire their data under very specific conditions (10 °C, 200 mM NaCl). The reason for this is that at under reasonable biological conditions (37 °C, 150 mM NaCl), their spectra showed evidence of an alternate conformation. At higher temperature and lower salt, the peaks corresponding to the structure at left disappeared entirely and were replaced by peaks corresponding to an alternate conformation.

You can see that alternate conformation, called Ltn40, at right (explore this structure at the PDB). As you can see, this structure is dimeric, taking the form of a β-sandwich. The β-sheets themselves have a sort of Greek key meander arrangement. You can learn more about β elements, protein topology, and protein structure at Larry Moran’s Sandwalk. This new fold buries a substantial number of hydrophobic side-chains between the β-sheets. At the same time, a preponderance of positively-charged residues is exposed to solution. The α-helices of the chemokine structure appear to unfold completely, a new β-strand forms at the N-terminus, and the existing sheets shift their hydrogen-bonding register by one residue, meaning that β1 and β3 are rotated 180° along their lengthwise axes.

The existence of two native folds under reasonable physiological conditions (Keq near one at 37 °C) poses two questions. The first is whether and how fast the structures interconvert. The fact that the equilibrium of a given sample can be shifted with temperature and ionic strength implies that this is an active equilibrium, i.e. that the energy barrier is low enough to cross using just thermal energy. Tuinstra et al. used an NMR experiment to establish that interconversion takes place on a timescale of about 100 ms.

The second question that naturally comes to mind is whether the two structures have different functions, and what those functions are. With a few clever experiments in which they used mutations to stabilize one fold or the other, Tuinstra et al. demonstrated that only the Ltn10 fold activates the partner GPCR, and only the Ltn40 fold binds to heparin, a polysaccharide often found in the ECM. Thus, neither fold possesses the full range of biological functions of lymphotactin; in order to fulfill its biological role it must switch between these structures in vivo. Moreover, the exclusivity of these functions between folds naturally suggests a switching mechanism for regulation.

This is an interesting and important finding because it is (so far) the only example of a protein adopting two completely different stable folds with no hydrogen bonds in common at equilibrium. Trivially, natively disordered proteins adopt multiple conformations under physiological solution conditions, and many proteins alter their conformations in response to ligand binding while keeping most of their hydrogen bond network intact. In this case, however, an existing network of stabilizing bonds is completely disrupted in order to form a new fold with a totally different function. I’ve already discussed some of the implications of this with respect to protein folding, and in regards to the recent transitive homology studies out of the Cordes group. Lymphotactin offers lessons and ideas for protein folding and evolution that must be taken into account. In particular, the fact that point mutations can significantly stabilize one or the other of these structures implies that there may be previously unsuspected shortcuts through structural space between folded states that avoid unproductive or energetically unfavorable molten globules.

In addition, these results signify that the Anfinsen paradigm that dominates our understanding of protein structure ought not be taken for granted. In many cases it is true that a peptide sequence uniquely determines a single structure under all physiological conditions. Of course we have known for some time that certain peptide sequences do not produce ordered structural ensembles at all. What the lymphotactin example makes crystal clear is that a given sequence can yield an ensemble with multiple energetic minima that reflect related but topologically distinct structures. Tuinstra et al. suspect that this phenomenon has not been noted previously because structures of this kind would not be amenable to crystallization, or would only crystallize in one (of many) structures. If this is so, then as more and more proteins are studied using solution techniques under physiological conditions we may find multiple structural minima in a variety of proteins. Such discoveries may significantly enhance our understanding of the protein regulation, function, and evolution.

1. Tuinstra, R.L., Peterson, F.C., Kutlesa, S., Elgin, E.S., Kron, M.A., Volkman, B.F. (2008). Interconversion between two unrelated protein folds in the lymphotactin native state. Proceedings of the National Academy of Sciences 105 (13) 5057-5062. DOI: 10.1073/pnas.0709518105

 Posted by at 3:10 AM