Proteins are the workhorses of the cell. The shape that a protein molecule adopts enables it to carry out its role. However, a protein’s shape, or 'conformation', is not static. Instead, a protein can shift between different conformations. This is particularly true for enzymes – the proteins that catalyze chemical reactions. The region of an enzyme where the chemical reaction happens, known as the active site, often has to change its conformation to allow catalysis to proceed. Changes in temperature can also make a protein shift between alternative conformations. Understanding how a protein shifts between conformations gives insight into how it works.

A common method for studying protein conformation is X-ray crystallography. This technique uses a beam of X-rays to figure out where the atoms of the protein are inside a crystal made of millions of copies of that protein. At room temperature or biological temperature, X-rays can rapidly damage the protein. Because of this, most crystal structures are determined at very low temperatures to minimize damage. But cooling to low temperatures changes the conformations that the protein adopts, and usually causes fewer conformations to be present.

Keedy, Kenner, Warkentin, Woldeyes et al. have used X-ray crystallography from a very low temperature (-173°C or 100 K) to above room temperature (up to 27°C or 300 K) to explore the alternative conformations of an enzyme called cyclophilin A. These alternative conformations include those that have previously been linked to this enzyme’s activity. Starting at a low temperature, parts of the enzyme were seen to shift from having a single conformation to many conformations above a threshold temperature. Unexpectedly, different parts of the enzyme have different threshold temperatures, suggesting that there isn’t a single transition across the whole protein. Instead, it appears the way a protein’s conformation changes in response to temperature is more complex than was previously realized. This result suggests that conformations in different parts of a protein are coupled to each other in complex ways.

Keedy, Kenner, Warkentin, Woldeyes et al. then performed X-ray crystallography at room temperature using an X-ray free-electron laser (XFEL). This technique can capture the protein’s structure before radiation damage occurs, and confirmed that the alternative conformations observed were not affected by radiation damage.

The combination of X-ray crystallography at multiple temperatures, new analysis methods for identifying and measuring alternative conformations, and XFEL crystallography should help future studies to characterize conformational changes in other proteins.

Current structural biology methods provide only incomplete pictures of how proteins interconvert between distinct conformations (Motlagh et al., 2014; van den Bedem and Fraser, 2015). Although X-ray crystallography reveals atomic coordinates with relatively high accuracy and precision, the resulting electron density maps contain contributions from multiple alternative conformations reflecting the ensemble average of 10⁶–10¹⁵ copies of the protein in one crystal (Rejto and Freer, 1996; Smith et al., 1986; Woldeyes et al., 2014). At high resolution, it is often possible to detect and discretely model these alternative conformations (Burnley et al., 2012; Davis et al., 2006; Lang et al., 2010; van den Bedem et al., 2009). Structural characterization of alternative conformations by X-ray crystallography can complement NMR (Baldwin and Kay, 2009; Fenwick et al., 2014) and computational simulations (Dror et al., 2012; Ollikainen et al., 2013) in defining the structural basis of protein dynamics and ultimately in linking dynamics to function (Henzler-Wildman and Kern, 2007).

However, more than 95% of crystal structures are determined at cryogenic temperatures (∼100 K) to reduce radiation damage by minimizing diffusion of reactive intermediates and chemical-damage-induced structural relaxations (Garman, 2010; Holton, 2009; Warkentin et al., 2013). Unfortunately, cryocooling can modify main chain and side chain conformational distributions throughout the protein, including at active sites and distal regions important for allosteric function (Fraser et al., 2011; Halle, 2004; Keedy, et al., 2014). Recent studies have instead used room temperature data collection to reveal a multitude of previously ‘hidden’ alternative conformations that are not evident at cryogenic temperatures, many of which have important ramifications for determining molecular mechanisms (Deis et al., 2014; Fraser et al., 2009; Fukuda and Inoue, 2015; van den Bedem et al., 2013).

Between these temperature extremes, protein conformational heterogeneity changes in complex ways. Previous studies using a wide variety of biophysical probes including NMR, X-ray crystallography, and neutron scattering have revealed a change in the character of conformational heterogeneity and/or protein dynamics around 180–220 K (Doster, 2010; Frauenfelder et al., 2009; Lewandowski et al., 2015; Ringe and Petsko, 2003) however, the molecular origins of this ‘glass’ or ‘dynamical’ transition remain incompletely understood. Classic work has examined the temperature dependence of protein conformational heterogeneity across individual X-ray structures determined at temperatures from ∼80 to 320 K (Frauenfelder et al., 1979, 1987; Tilton et al., 1992). These studies used atomic B-factors as a proxy for conformational heterogeneity and identified a global inflection point around 180–220 K. This inflection point was interpreted in terms of a transition driven by dynamical arrest of the coupled hydration layer-protein system (Doster et al., 1989; Frauenfelder et al., 1979, 1987; Tilton et al., 1992). By contrast, solution NMR studies of picosecond–nanosecond (ps–ns) timescale methyl side chain order parameters showed heterogeneous changes in motional amplitudes at temperatures between 288 and 346 K. Thermal deactivation of these motions was suggested to predict a transition near 200 K without invoking solvent arrest (Lee and Wand, 2001). Recent solid-state NMR (ssNMR) experiments suggest that protein motions are coupled to solvent, and that three transitions at ∼195, 220, and 250 K mark the onset of distinct classes of motions as temperature increases (Lewandowski et al., 2015). Unfortunately, these studies used either globally averaged data (as with ssNMR or neutron scattering) or imprecise atomic-level models of conformational heterogeneity (as with B-factors in X-ray crystallography or NMR order parameters), thus preventing an all-atom understanding of the complex temperature response of protein crystals.

New crystallographic and computational techniques now enable a more detailed investigation of the temperature dependence of protein conformational heterogeneity at the atomic level. First, the program Ringer (Lang et al., 2014, 2010) evaluates low-level electron density traditionally considered noise to uncover statistically significant ‘hidden’ alternative conformations, which may become populated or depopulated as a function of temperature. Second, multiconformer models with explicit alternative conformations of both backbone and side chain atoms, as created by manual building or methods such as the program qFit (Keedy et al., 2015, van den Bedem et al., 2009), can account for non-harmonic motions across separate energy wells (encoded by discrete alternative conformations with distinct occupancies and coordinates) and harmonic motions within energy wells (encoded by B-factors). Third, crystallographic order parameters (S²) weigh these harmonic and non-harmonic contributions in a single metric that quantifies the disorder of each residue in a multiconformer model, allowing direct comparison with NMR-determined order parameters (Fenwick et al., 2014). Finally, methodological advances based on the physics of ice formation have enabled variable-temperature crystallographic data collection at temperatures between 300 and 100 K with modest or no use of potentially conformation-perturbing cryoprotectants (Warkentin et al., 2012; Warkentin and Thorne, 2009). Together, these methods overcome many of the limitations of previous X-ray-based approaches and will contribute to an integrated view of how protein conformational heterogeneity and dynamics evolve with temperature.

The human proline isomerase cyclophilin A (CypA) is an excellent model system for deploying these tools to study the structural basis of functional conformational dynamics and, in particular, to use temperature to understand the extent of correlated motions during an enzyme’s catalytic cycle. Previous NMR relaxation data for CypA (Eisenmesser et al., 2005, 2002) indicated a single common exchange process, both in the apo state and during catalysis, for a network of dynamic residues extending from the core to the active site. Room temperature crystallography later suggested the precise alternative conformations that collectively interconvert during catalysis (Fraser et al., 2009). However, subsequent NMR relaxation experiments of mutants designed to perturb the dynamics suggested that multiple exchange processes occur within this network (Schlegel et al., 2009). Here, we analyze multitemperature synchrotron experiments to examine the temperature-dependent conformational heterogeneity of CypA. Additionally, we report X-ray-free electron laser (XFEL) data, which are free of conventional radiation damage (Kern et al., 2014; Spence et al., 2012), to validate previous connections between alternative conformations determined by synchrotron crystallography and NMR experiments performed in solution (Eisenmesser et al., 2005; Fraser et al., 2009). Our analysis shows that the temperature dependence of alternative protein conformations is heterogeneous and that the character of this heterogeneity bridges previous models for protein dynamical transitions. Our results also suggest new ways to use variable temperature with both synchrotron and XFEL crystallography to probe the dynamic underpinnings of protein function.

Here, we have mapped the conformational landscape of the dynamic enzyme CypA by analyzing multiconformer models from multitemperature crystallography. Unlike previous temperature-dependent analyses of X-ray crystallography, here we consider both harmonic disorder (B-factors) and non-harmonic displacements (alternative conformations and occupancies), characterized using crystallographic bond order parameters. We have four primary findings:

1. The occupancy of alternative side chain conformations present at 310 K generally decreases with decreasing temperature.

2. However, some residues adopt alternative conformations only at 180 K and below.

3. The temperature response of residues is highly heterogeneous: the ‘intersection temperature’ below which side chain conformational heterogeneity remains nearly constant has a broad distribution across the molecule, with a peak near 250 K.

4. Residues composing a dynamic active-site network that is important for function do not have a consistent intersection temperature.

Our results provide new insight into the relationship between energy landscapes, the glass transition, and protein function. Glasses and other disordered systems have complex energy landscapes, in part due to the large number of microenvironments and the extensive frustration that disorder generates in the intermolecular interactions. Proteins at biological temperatures are ‘glassy’ in this sense—they have complex energy landscapes, due to their large size, heterogeneous amino acid composition, and many degrees of conformational freedom (Frauenfelder et al., 1991). The extensive heterogeneity in the temperature response of individual residues in CypA that we observe here provides additional direct evidence for this underlying energetic heterogeneity.

In addition to the inherent ‘glassiness’ of proteins at biological temperatures, dynamical transitions, some of which have been called ‘glass transitions’, have been reported at lower temperatures, including 180, 200, 220, 240, and 250 K, based on Mössbauer spectroscopy, X-ray crystallography, liquid and solid-state NMR, neutron scattering, and other techniques (Lewandowski et al., 2015; Schiro et al., 2015). These transitions typically manifest as a change in slope of some measurement in the vicinity of the suggested transition temperature. However, many of these measurements are sensitive to motions only within some timescale window (often ps-ns), monitor only a subset of amino acid types, and/or spatially average over all residues in the protein. By contrast, multitemperature crystallography with multiconformer models has many advantages by providing a time-independent and fully site-resolved measurement of ensemble-averaged atomic displacements, including both harmonic and discrete conformational heterogeneity, within the crystal.

This combined methodology lets us examine dynamical and glass transitions in protein crystals from a new perspective. Glass transitions are by definition non-equilibrium phenomena that arise when the kinetics of relaxation toward equilibrium slow so dramatically that equilibrium cannot be reached on experimental timescales. One signature of a true glass transition in proteins would be if occupancies of minority alternative conformations were arrested at non-zero values below some temperature. Indeed, here we see no appreciable temperature evolution of individual conformer occupancies or B-factors at 180 K and below, and the average disorder at these temperatures is far in excess of what is predicted from high-temperature extrapolations. These observations are consistent with the falling out of equilibrium expected in a glass transition, but not with a transition driven by the thermal freeze-out of alternative side chain conformations (Lee and Wand, 2001). Moreover, the persistence of multiple rotameric states at low temperatures is consistent with solvent arrest that impedes further changes in side chain disorder. Although the details of protein–solvent interactions may differ in crystals versus in solution, local variability at different protein–solvent interface microenvironments in the crystal (Teeter et al., 2001) likely contributes to the heterogeneity of temperature responses that we observe. The critical importance of site resolution is evident in the results of Figure 6. Averaging over side chain disorder in all residues yields an apparent transition near 250 K (Figure 6 and Figure 6—figure supplement 7). Perhaps coincidentally, this same temperature has been associated with a transition for protein side chains in site-averaged solid-state NMR measurements (Lewandowski et al., 2015). However, the ‘transition’ in our residue-averaged result obscures the highly heterogeneous temperature dependence of the individual side chains. We find no evidence of tight cooperativity or of a collective global response near 250 K that would be expected in the case of a ‘true’ dynamical transition. Instead, our data are consistent with local, non-cooperative freeze-out of conformational states defined by the energy landscape over a broad temperature range.

The heterogeneous response of side chain order parameters to temperature is driven largely by the changes to the populations of alternative conformations, which ‘flat-line’ at different temperatures across CypA. This diversity of ‘flat-lining’ temperatures is present even within the dynamic active-site network, even though the constituent residues have similar occupancies for their major and minor states at high temperatures (Figure 7). This result contrasts with previous NMR and X-ray experiments that hypothesized correlated motions of this network as rate-limiting for the catalytic cycle (Eisenmesser et al., 2005; Fraser et al., 2009) (Figure 8A).

Figure 8

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To bridge these views, we propose that the active-site network adopts two substates, which are primarily distinguished by Phe113 rotamer interconversion. Each of these substates adopts a differently weighted ensemble of conformations for other residues (Figure 8B). In this model, Met61 and Arg55 can switch rotamers more easily than Ser99 when Phe113 is in its χ1 gauche (p) rotamer pointed toward Ser99, whereas Ser99 can switch rotamers more easily than Met61 and Arg55 when Phe113 is in its χ1 gauche (m) rotamer pointed toward Met61. Additionally, thermal ‘breathing’ motions within rotameric wells may relieve minor steric overlaps within some of these macro- and microstates (Figure 8B , bottom right). This model is consistent with Phe113 having the lowest ‘flat-lining’ temperature of the network (Figure 7), and makes sense sterically because of the large size of the aromatic ring. These hypothesized motions are consistent with the timescales and temperature dependencies of motion assigned by solid-state NMR studies of crystalline protein GB1 (Lewandowski et al., 2015). Furthermore, the model helps explain the difficulty of fitting NMR relaxation data for perturbed versions of the active-site network as a single collective exchange process (Schlegel et al., 2009). The aromatic ring of Phe113 could play a dominant role in determining the chemical shift changes of the surrounding residues. Each of these residues could also populate multiple rotamers in the excited state measured by NMR. Our hierarchical perspective evokes the ‘population shuffling’ model of (Smith et al., 2015), in which a protein macrostate (in CypA, defined by the Phe113 rotamer) also determines the different relative populations of rotamers for a subset of other residues (in CypA, the other residues in the active-site network). In this model, the interconversion between macrostates, and not the collective motion of all residues between distinct rotamers, is correlated with the rate-limiting step of the CypA catalytic cycle.

Diversity in the temperature dependences of alternative conformations as we see here is inevitable given the limitations of the amino acid alphabet, yet its spatial pattern within a protein may provide insight into selective pressures. Evolutionary optimization must ensure that functionally important alternative conformations are robustly populated and interconvert appreciably at physiological conditions. However, the energy landscapes of individual residues are coupled to varying extents, such that some subsets of residues must be collectively optimized to preserve some, but not perfect, collectivity in functional motions. For proteins with large sequence alignments, evolutionary covariation has been used to predict ‘sectors’ of functionally cooperative residues, which are often dispersed in primary sequence but strikingly contiguous in tertiary structure (Halabi et al., 2009). By contrast, temperature-dependent crystallography has the potential to unveil couplings in atomic detail by identifying sets of residues whose conformational ensembles respond concertedly to temperature change. Based on our results with CypA, we expect this coupling to be weak, but measurable. Serial femtosecond XFEL crystallography combined with ultra-fast temperature jumps could enable a temporal view of these coupled conformational changes. Novel static and time-resolved multitemperature crystallographic approaches will provide powerful tools for resolving concerted motions to explore how proteins function and evolve.

1. 1. Keedy DA
   2. Kenner LR
   3. Warkentin M
   4. Woldeyes RA
   5. Thompson MC
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   7. Van Benschoten AH
   8. Baxter EL
   9. Hopkins JB
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   11. McPhillips SE
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   17. Soltis M
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   19. Gonzalez A
   20. Sauter NK
   21. Cohen AE
   22. van den Bedem H
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   24. Fraser JS

   (2015) Multiconformer synchrotron model of CypA at 100 K

   Publicly available at the RCSB Protein Data Bank (Accession no: 4yug).

   http://www.rcsb.org/pdb/explore/explore.do?structureId=4YUG
2. 1. Keedy DA
   2. Kenner LR
   3. Warkentin M
   4. Woldeyes RA
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   9. Hopkins JB
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   24. Fraser JS

   (2015) Multiconformer synchrotron model of CypA at 150 K

   Publicly available at the RCSB Protein Data Bank (Accession no: 4yuh).

   http://www.rcsb.org/pdb/explore/explore.do?structureId=4YUH
3. 1. Keedy DA
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   (2015) Multiconformer synchrotron model of CypA at 180 K

   Publicly available at the RCSB Protein Data Bank (Accession no: 4yui).

   http://www.rcsb.org/pdb/explore/explore.do?structureId=4YUI
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   (2015) Multiconformer synchrotron model of CypA at 240 K

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   http://www.rcsb.org/pdb/explore/explore.do?structureId=4YUJ
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   Publicly available at the RCSB Protein Data Bank (Accession no: 4yuk).

   http://www.rcsb.org/pdb/explore/explore.do?structureId=4YUK
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   http://www.rcsb.org/pdb/explore/explore.do?structureId=4YUL
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   http://www.rcsb.org/pdb/explore/explore.do?structureId=4YUM
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   http://www.rcsb.org/pdb/explore/explore.do?structureId=4YUN
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Author details

1. Daniel A Keedy

   Department of Bioengineering and Therapeutic Sciences, University of California, San Francisco, San Francisco, United States

   Contribution

   DAK, Conception and design, Analysis and interpretation of data, Drafting or revising the article

   Contributed equally with

   Lillian R Kenner, Matthew Warkentin and Rahel A Woldeyes

   Competing interests

   No competing interests declared.

   "This ORCID iD identifies the author of this article:" 0000-0002-9184-7586

2. Lillian R Kenner

   Department of Bioengineering and Therapeutic Sciences, University of California, San Francisco, San Francisco, United States

   Contribution

   LRK, Conception and design, Acquisition of data, Drafting or revising the article

   Contributed equally with

   Daniel A Keedy, Matthew Warkentin and Rahel A Woldeyes

   Competing interests

   No competing interests declared.

3. Matthew Warkentin

   Department of Physics, Cornell University, Ithaca, United States

   Contribution

   MW, Conception and design, Acquisition of data, Drafting or revising the article

   Contributed equally with

   Daniel A Keedy, Lillian R Kenner and Rahel A Woldeyes

   Competing interests

   No competing interests declared.

4. Rahel A Woldeyes

   Department of Bioengineering and Therapeutic Sciences, University of California, San Francisco, San Francisco, United States

   Contribution

   RAW, Conception and design, Acquisition of data, Analysis and interpretation of data, Drafting or revising the article

   Contributed equally with

   Daniel A Keedy, Lillian R Kenner and Matthew Warkentin

   Competing interests

   No competing interests declared.

   "This ORCID iD identifies the author of this article:" 0000-0003-0737-8383

5. Jesse B Hopkins

   Department of Physics, Cornell University, Ithaca, United States

   Contribution

   JBH, Acquisition of data, Drafting or revising the article

   Competing interests

   No competing interests declared.

6. Michael C Thompson

   Department of Bioengineering and Therapeutic Sciences, University of California, San Francisco, San Francisco, United States

   Contribution

   MCT, Analysis and interpretation of data, Drafting or revising the article

   Competing interests

   No competing interests declared.

7. Aaron S Brewster

   Physical Biosciences Division, Lawrence Berkeley National Laboratory, Berkeley, United States

   Contribution

   ASB, Acquisition of data, Analysis and interpretation of data, Drafting or revising the article

   Competing interests

   No competing interests declared.

8. Andrew H Van Benschoten

   Department of Bioengineering and Therapeutic Sciences, University of California, San Francisco, San Francisco, United States

   Contribution

   AHVB, Acquisition of data, Analysis and interpretation of data, Drafting or revising the article

   Competing interests

   No competing interests declared.

9. Elizabeth L Baxter

   Stanford Synchrotron Radiation Lightsource, SLAC National Accelerator Laboratory, Menlo Park, United States

   Contribution

   ELB, Acquisition of data, Drafting or revising the article

   Competing interests

   No competing interests declared.

10. Monarin Uervirojnangkoorn

    1. Department of Molecular and Cellular Physiology, Stanford University, Stanford, United States
    2. Howard Hughes Medical Institute, Stanford University, Stanford, United States

    Contribution

    MU, Analysis and interpretation of data, Drafting or revising the article, Contributed unpublished essential data or reagents

    Competing interests

    No competing interests declared.

11. Scott E McPhillips

    Stanford Synchrotron Radiation Lightsource, SLAC National Accelerator Laboratory, Menlo Park, United States

    Contribution

    SEM, Acquisition of data, Drafting or revising the article

    Competing interests

    No competing interests declared.

12. Jinhu Song

    Stanford Synchrotron Radiation Lightsource, SLAC National Accelerator Laboratory, Menlo Park, United States

    Contribution

    JS, Acquisition of data, Drafting or revising the article

    Competing interests

    No competing interests declared.

13. Roberto Alonso-Mori

    Linac Coherent Light Source, SLAC National Accelerator Laboratory, Menlo Park, United States

    Contribution

    RA-M, Acquisition of data, Drafting or revising the article

    Competing interests

    No competing interests declared.

14. James M Holton

    1. Physical Biosciences Division, Lawrence Berkeley National Laboratory, Berkeley, United States
    2. Stanford Synchrotron Radiation Lightsource, SLAC National Accelerator Laboratory, Menlo Park, United States
    3. Department of Biochemistry and Biophysics, University of California, San Francisco, San Francisco, United States

    Contribution

    JMH, Analysis and interpretation of data, Drafting or revising the article

    Competing interests

    No competing interests declared.

15. William I Weis

    1. Department of Molecular and Cellular Physiology, Stanford University, Stanford, United States
    2. Department of Structural Biology, Stanford University, Stanford, United States
    3. Department of Photon Science, SLAC National Accelerator Laboratory, Menlo Park, United States

    Contribution

    WIW, Analysis and interpretation of data, Drafting or revising the article, Contributed unpublished essential data or reagents

    Competing interests

    No competing interests declared.

    "This ORCID iD identifies the author of this article:" 0000-0002-5583-6150

16. Axel T Brunger

    1. Department of Molecular and Cellular Physiology, Stanford University, Stanford, United States
    2. Howard Hughes Medical Institute, Stanford University, Stanford, United States
    3. Department of Structural Biology, Stanford University, Stanford, United States
    4. Department of Photon Science, SLAC National Accelerator Laboratory, Menlo Park, United States

    Contribution

    ATB, Acquisition of data, Drafting or revising the article, Contributed unpublished essential data or reagents

    Competing interests

    ATB: Reviewing editor, eLife.

    "This ORCID iD identifies the author of this article:" 0000-0001-5121-2036

17. S Michael Soltis

    Stanford Synchrotron Radiation Lightsource, SLAC National Accelerator Laboratory, Menlo Park, United States

    Contribution

    SMS, Acquisition of data, Drafting or revising the article

    Competing interests

    No competing interests declared.

    "This ORCID iD identifies the author of this article:" 0000-0003-4678-2995

18. Henrik Lemke

    Linac Coherent Light Source, SLAC National Accelerator Laboratory, Menlo Park, United States

    Contribution

    HL, Acquisition of data, Drafting or revising the article

    Competing interests

    No competing interests declared.

19. Ana Gonzalez

    Stanford Synchrotron Radiation Lightsource, SLAC National Accelerator Laboratory, Menlo Park, United States

    Contribution

    AG, Acquisition of data, Drafting or revising the article

    Competing interests

    No competing interests declared.

20. Nicholas K Sauter

    Physical Biosciences Division, Lawrence Berkeley National Laboratory, Berkeley, United States

    Contribution

    NKS, Analysis and interpretation of data, Drafting or revising the article

    Competing interests

    No competing interests declared.

21. Aina E Cohen

    Stanford Synchrotron Radiation Lightsource, SLAC National Accelerator Laboratory, Menlo Park, United States

    Contribution

    AEC, Acquisition of data, Drafting or revising the article

    Competing interests

    No competing interests declared.

22. Henry van den Bedem

    Stanford Synchrotron Radiation Lightsource, SLAC National Accelerator Laboratory, Menlo Park, United States

    Contribution

    HvdB, Conception and design, Acquisition of data, Analysis and interpretation of data, Drafting or revising the article

    For correspondence

    vdbedem@slac.stanford.edu

    Competing interests

    No competing interests declared.

    "This ORCID iD identifies the author of this article:" 0000-0003-2358-841X

23. Robert E Thorne

    Department of Physics, Cornell University, Ithaca, United States

    Contribution

    RET, Conception and design, Analysis and interpretation of data, Drafting or revising the article

    For correspondence

    ret6@cornell.edu

    Competing interests

    No competing interests declared.

24. James S Fraser

    Department of Bioengineering and Therapeutic Sciences, University of California, San Francisco, San Francisco, United States

    Contribution

    JSF, Conception and design, Acquisition of data, Analysis and interpretation of data, Drafting or revising the article

    For correspondence

    jfraser@fraserlab.com

    Competing interests

    No competing interests declared.

    "This ORCID iD identifies the author of this article:" 0000-0002-5080-2859

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