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Structural and functional characterization of dynamic oligomerization in Burkholderia cenocepacia HMG-CoA reductase Riley B. Peacock, Chad W. Hicks, Alexander M. Walker, Sophia M. Dewing, Kevin M. Lewis, Jean-Claude Abboud, Samuel W. A. Stewart, ChulHee Kang, and Jeffrey Michael Watson Biochemistry, Just Accepted Manuscript • DOI: 10.1021/acs.biochem.9b00494 • Publication Date (Web): 30 Aug 2019 Downloaded from pubs.acs.org on August 30, 2019

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Biochemistry

Structural and functional characterization of dynamic oligomerization in Burkholderia cenocepacia HMG-CoA reductase Riley B. Peacock†1, Chad W. Hicks†2, Alexander M. Walker‡, Sophia M. Dewing†, Kevin M. Lewis‡, Jean-Claude Abboud†, Samuel W. A. Stewart†, ChulHee Kang‡, Jeffrey M. Watson†* †Department

of Chemistry and Biochemistry, Gonzaga University, Spokane, Washington, 99258,

United States and ‡Department of Chemistry, Washington State University, Pullman, Washington, 99164, United States

ACS Paragon Plus Environment

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ABSTRACT

The enzyme 3-hydroxy-3-methylglutaryl coenzyme A (HMG-CoA) reductase (HMGR), in most organisms, catalyzes the four-electron reduction of the thioester (S)-HMG-CoA to the primary alcohol (R)-mevalonate, utilizing NADPH as the hydride donor. In some organisms, including the opportunistic lung pathogen Burkholderia cenocepacia, it catalyzes the reverse reaction, utilizing NAD+ as a hydride acceptor in the oxidation of mevalonate. Burkholderia cenocepacia HMGR has been previously shown to exist as an ensemble of multiple non-additive oligomeric states, each with different levels of enzymatic activity, suggesting that the enzyme exhibits characteristics of the morpheein model of allostery. We have characterized a number of factors, including pH, substrate concentration and enzyme concentration, that modulate the structural transitions that influence the interconversion among the multiple oligomers. We have also determined the crystal structure of Burkholderia cenocepacia HMGR in the hexameric state bound to coenzyme A and ADP. This hexameric assembly provides important clues as to how the transition among oligomers might occur, and why Burkholderia cenocepacia HMGR, unique among characterized HMGRs, exhibits morpheein-like behavior. INTRODUCTION Enzymatic activity is regulated by a wide range of mechanisms, including post-translational modification, transcriptional control, regulated proteolysis, allostery and many more. This diversity of tools allows a cell to finely tune the rate of an enzyme-catalyzed reaction to respond to changes in the environment, whether intracellular or extracellular. Allostery, in particular, provides a mechanism by which local cellular concentrations of metabolites can influence the rate of key reactions. Since the discovery of allosteric regulation of binding proteins and


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Biochemistry

enzymes over 100 years ago, the structural mechanisms by which these changes in activity occur have been an active area of study. As a consequence, several models of allostery have been proposed to describe how a protein’s structure changes from the more active state (often called the relaxed, or R, state) to the less active state (often called the tense, or T, state). For example, the concerted (MWC) model is characterized by large rearrangements of the quaternary structure, resulting in a protein that is either all R- or all T-state (1). In this model, changes in concentration of allosteric effectors influence the equilibrium constant between the two states. The sequential (KNF) model, on the other hand, is generally characterized by smaller changes in tertiary structure, resulting in a protein that can range from all R-state, to partly R- and partly Tstate, to all T-state(2). In this model, changes in the concentration of effectors modulate the degree to which the protein is in the R-state or T-state. In both models, however, the overall level of quaternary structure generally does not change, or modulates between additive quaternary structures. For example, the canonical allosteric protein hemoglobin, which exhibits characteristics of both models, remains a tetramer regardless of whether it is in the R-state or Tstate (reviewed, for example, in (3)). Phosphofructokinase-1, which catalyzes the first committed step in glycolysis, changes between an inactive (T-state) dimer and an active (R-state) tetramer, or dimer of dimers, in response to some allosteric effectors (4). A recently proposed model of allostery differs in some principal assumptions when compared to the first two models. The morpheein model of allostery (5) describes a protein existing as a dynamic ensemble of at least two different, non-additive quaternary forms, whose interconversion requires multimer dissociation. Alternate oligomers have alternate functions, which for an enzyme can be different levels of activity. Because these forms are non-additive, there can be no direct equilibrium between them. For example, the first morpheein identified,


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porphobilinogen synthase, takes on both hexameric and octameric quaternary structures, and both can be present at the same time (6, 7). Transition between states proceeds through a dissociation to a lower order form, followed by a conformational change, ending with a reassociation of the new structure into an alternate quaternary form. Substrates and other effectors differentially stabilize one or the other assembly, and the enzymatic activity increases or decreases as a result. In addition, changes in pH and enzyme concentration are also known to modulate the quaternary structure transition in morpheeins, due to their effects on protein-protein interactions and the law of mass action (8). The family of proteins that have been identified as morpheeins or likely morpheeins remains relatively small (9-12). One such protein is the 3-hydroxy-3-methylglutaryl coenzyme A (HMGCoA) reductase from the opportunistic lung pathogen Burkholderia cenocepacia (13). Like other HMG-CoA reductases, Bc HMG-CoA reductase (BcHMGR) catalyzes the reversible fourelectron reduction of the thioester HMG-CoA to the primary alcohol mevalonate (Figure 1), using either NAD(H) or NADP(H) as its hydride carrier, depending on the species from which it is isolated.

NAD(P)+

O H C OH O 3 SCoA

O

NAD(P)H

(S)-HMG-CoA

O H C OH OH 3 O

SCoA

O H C OH O 3

CoASH

H

O

O H C OH OH 3

NAD(P)+

H

O NAD(P)H

[mevaldyl-CoA]

[mevaldehyde]

(R)-mevalonate

Figure 1. Reaction catalyzed by HMG-CoA reductase. The four-electron reduction, as written, proceeds through two putative intermediates, neither of which is released over the course of the reaction. Some bacterial HMG-CoA reductases, including that from Burkholderia cenocepacia, likely utilize the reverse, oxidative reaction in vivo.


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BcHMGR, like the well-characterized HMGR from Pseudomonas mevalonii (PmHMGR) (14), strongly favors NAD(H) over NADP(H) as its hydride carrier, suggesting that it likely works in the oxidative (mevalonate to HMG-CoA) direction in the cell. Unlike other HMGRs, however, BcHMGR was shown to exist as an ensemble of three different quaternary structures, initially characterized by size-exclusion chromatography as a hexamer, a nonamer, and a larger assembly initially designated as an 18-mer (13) . In addition, initial kinetic characterization showed that for each of the three substrates of the oxidative reaction – mevalonate, NAD+ and coenzyme A – BcHMGR demonstrates double-saturation kinetics characteristic of PBGS, the prototype morpheein, suggesting that these oligomers each exhibited different Vmax and KM values. These lines of evidence suggested that BcHMGR was likely a morpheein, though a significant amount of characterization remained in order to understand the factors that might modulate the transition between the individual structural states (13). We sought to further explore these factors and further verify the suspected morpheein behavior that may impart a complex regulatory scheme on BcHMGR that is not echoed in closely related HMGRs. In the following work, we demonstrate that pH and enzyme concentration have dramatic effects on both the kinetics and activity level of the enzyme, as well as on the distribution of quaternary forms in the ensemble. We also show that changes in substrate concentration result in structural change as judged by changes in solution exposure of hydrophobic area. Finally, we have also determined the crystal structure of the hexameric state of BcHMGR in complex with coenzyme A and ADP. This allows us to correlate BcHMGR solution behavior with structural evidence, and develop preliminary theories for why BcHMGR exhibits morpheein-like behavior as well as why other closely related HMGRs might not share this intricate interplay of dynamic oligomerization and activity change.


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MATERIALS AND METHODS Protein Expression and Purification Burkholderia cenocepacia HMG-CoA reductase (UniProtKB accession ID: B4EKH5) was expressed and purified as previously described (13). Buffer species used were as follows: 20 mM HEPES (pH 7.5), 20 mM tricine (pH 8.3) or 20 mM glycine (pH 9.0). All buffers also included 50 mM KCl. Protein samples were concentrated, where appropriate, using Vivaspin concentrators (Viva Products) with a 30K molecular weight cutoff. Protein concentration was measured by absorbance at 280 nm using the previously established extinction coefficient of 22,920 M-1 cm-1 (13) . Enzyme Kinetics BcHMGR was assayed according to previously published procedures (13). In summary, an assay mixture contained buffer of appropriate pH, NAD+, mevalonate, and BcHMGR of appropriate concentration. Reactions were initiated by addition of coenzyme A and absorbance was monitored at 340 nm for 1 minute. Rates were determined from the slope of the best-fit line to the linear initial rate portion of the progress curve. For measurement of kcat, a typical assay mixture contained 5.94 mM mevalonate, 3.96 mM NAD+, 5.98 mM coenzyme A and enzyme in buffer of appropriate pH. These represent saturating substrate concentrations at pH 7.5 and therefore Vmax conditions. These substrate concentrations were used for assays performed at all pH values. For determination of reaction rate as a function of coenzyme A concentrations at varying values of pH, mevalonate and NAD+ were held constant at the same concentrations as


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above, while coenzyme A was varied. All assays were performed in triplicate in a final volume of 500 L at 37 °C on a Cary 50 UV-Vis spectrophotometer (Agilent Technologies) equipped with a single-cell Peltier thermal controller. Size Exclusion Chromatography Size exclusion chromatography was performed using a HiLoad 16/60 Superdex 200 prep grade column (GE Healthcare) equilibrated with buffer of appropriate pH as listed above. A standard curve was generated using a Gel Filtration Markers Kit for protein molecular weights 29 to 700 kD (Sigma-Aldrich) prepared according to standard protocol. Column void volume was determined using Blue Dextran (Sigma-Aldrich). 4 mL of BcHMGR was injected onto the column at known concentration and eluted with the same buffer used for equilibration at a flow rate of 0.5 mL/min. Elution progress was monitored by absorbance at 280 nm. Results are the average of triplicate trials. ANS Fluorescence Assays were performed in 20 mM HEPES, pH 7.5, 50 mM KCl buffer, with a final 8-anilino-1naphthalenesulfonic acid (ANS) concentration of 30 M. BcHMGR was added to final concentrations of 0.5, 1.0 or 2.0 mg/mL as appropriate to an initial assay volume of 2 mL and incubated for 1 minute. Substrates (coenzyme A, NAD+, or NADH) were titrated into the samples and incubated for 1 minute. Each sample was excited at 370 nm, with emission monitored from 450-490 nm in a Cary Eclipse spectrofluorimeter (Agilent Technologies). Emission intensities at 470 nm were collected and corrected for dilution. Final assay volumes for each sample were 2.2 mL, indicating initial concentrations of protein and ANS changed by


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no more than 10% over the course of the assay. Results are the average of triplicate trials, presented as percent of initial ANS fluorescence in the absence of substrate. Crystallization Burkholderia cenocepacia HMG-CoA reductase (7.0 mg/mL, 20 mM HEPES, pH 7.5, 50 mM KCl) incubated overnight with 10 mM coenzyme A crystallized in a 1:1 mixture of protein and screening solution (0.2 M potassium acetate, pH 7.5, 20% (w/v) PEG 3,350). Crystals were grown by hanging drop vapor diffusion using a Phoenix RE (Art Robbins Instruments) robot and incubated at 4 °C. Crystals generally appeared after three days. Structure Determination Crystallographic data were collected at the Advanced Light Source (Beamline 8.2.1) and were reduced and scaled using HKL2000 (15). The structure of Burkholderia cenocepacia HMGCoA reductase was solved by molecular replacement in PHENIX (16) with the structure of Delftia acidovorans HMG-CoA reductase (PDB ID 6DIO) serving as the input model. Iterative model adjustment and refinement were completed using COOT (17)and PHENIX. Data collection and refinement statistics are provided in Table 1. Crystallographic coordinates and structure factors for Burkholderia cenocepacia HMG-CoA reductase have been deposited in the Protein Data Bank as PDB ID 6P7K.


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Table 1. X-ray data collection and refinement statisticsa HMG-CoA reductase Data collection Space group P6322 Cell dimensions a, b, c (Å) 141.489, 141.489, 122.969 α, β, γ (°) 90, 90, 120 Resolution (Å) 43.4 – 1.741 (1.809 – 1.747) Rmerge 0.1201 (1.095) Wavelength (Å) 1 Unique reflections 73523 (7175) Completeness (%) 89.07 (73.96) /σI 15.19 (2.02) CC1/2 0.997 (0.798) Redundancy 19.9 (10.7) Refinement Rwork / Rfree 0.1685 / 0.1898 (0.2829 / 0.3355) Number of atoms Protein and ligand 2852 Water 423 2 B-factors (Å ) All atoms 28.35 Solvent 40.89 Ligands 44.08 (ADP), 33.87 (CoA) R.m.s deviations Bonds (Å) 0.015 Angles (º) 1.69 Ramachandrans % Favored 97.05 % Outliers 0 Clashscore 1.93 a Statistics for the highest-resolution shell are shown in parentheses. Molecular Dynamics Simulations Molecular dynamics simulations were performed with NAMD 2.12 (18). Two dimers of Burkholderia cenocepacia HMG-CoA reductase identified via the PISA server (19) as interacting via the N-terminal coenzyme A binding motif were used as the initial model. After


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bathing the initial model in a solvent sphere, simulations were performed using a 2 fs time step with temperature maintained at 300 K using Langevin dynamics with a damping coefficient of 1/ps. The CHARMM36 force field (20) and TIP3P (21) water model were used in all simulations. RESULTS pH and Enzyme Concentration Dramatically Affect the Kinetic Properties of BcHMGR As demonstrated previously, the presence of multiple active oligomeric states of BcHMGR at pH 7.5 results in a double-saturation Vo vs. [S] plot (13). The kinetics of the oxidative BcHMGR reaction were studied at pH 7.5, 8.3 and 9.0, with mevalonate and NAD+ held constant at saturating concentrations and varying CoA. The shape of the Vo vs. [S] plot changes as pH increases, shifting from the double-saturation shape at 7.5 (Figure 2; green) and 8.3 (Figure 2; blue) to a nearly hyperbolic shape at pH 9.0 (Figure 2; red), suggesting that the distribution of oligomers strongly favors a single state. At pH 9.0, BcHMGR shows a Hill coefficient of 1.46 with a K0.5 of 0.061 mM. The only other HMGR shown to exhibit positive cooperativity, from Listeria monocytogenes, has Hill coefficients of 2.20 and 2.21 for the reductive reaction substrates HMG-CoA and NADPH respectively, but little to no cooperativity toward the oxidative substrates (22).


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Figure 2. BcHMGR activity as a function of pH. In all three trials, specific activity is given in mol NAD+ oxidized per minute per mg of enzyme. Mevalonate was held constant at 6 mM, NAD+ at 4 mM and coenzyme A varied. BcHMGR was held at 0.5 mg/mL in all trials. Points represent the average of triplicate trials, with error bars representing one standard deviation. Where error bars are not visible, they are smaller than the width of the symbol. pH values are 7.5 (green), 8.3 (blue), and 9.0 (red).

Earlier kinetic studies of BcHMGR showed that each of the oligomeric states exhibited a different Vmax, typical of a morpheein enzyme. As seen in Figure 2, the maximum specific activity also depends on pH. The highest specific activity (~11 mol NAD+ reduced/min/mg of enzyme) is achieved at pH 8.3, nearly twice that of pH 7.5 (~6 mol NAD+ reduced/min/mg enzyme) or pH 9.0 (5.89±0.22 mol NAD+ reduced/min/mg of enzyme), suggesting that a higher activity state might be more prevalent at this pH than at the other two. All three experiments were performed at the same enzyme concentration (0.5 mg/mL), so this difference in maximal activity must arise from a change in kcat as the distribution of oligomers change. This behavior has been observed for other morpheeins and is considered diagnostic for morpheein behavior (8). To test this hypothesis, kinetic assays were performed under saturating substrate concentrations


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at several enzyme concentrations. There is a clear effect of enzyme concentration on the kcat value for the enzyme-catalyzed reaction (Figure 3). As enzyme concentration increases, kcat initially increases as well, but exhibits a steady decline as enzyme concentration continues to increase. The oligomer with the highest activity appears to be favored at lower enzyme concentrations at pH 7.5.

Figure 3. Effect of enzyme concentration on kcat at pH 7.5. Mevalonate and coenzyme A were held constant at 6 mM, NAD+ was held constant at 4 mM, representing saturating substrate conditions. Points represent the average of triplicate trials, with error bars representing one standard deviation. Where error bars are not visible, they are smaller than the width of the symbol.

Relative Distribution of Quaternary States of BcHMGR Changes With pH and Enzyme Concentration Given the evident changes in kinetic behavior as a function of solution pH and enzyme concentration, size exclusion chromatography (SEC) was utilized to study the oligomeric states


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Biochemistry

of BcHMGR at different values of pH and enzyme concentration. As stated earlier, initial SEC characterization at pH 7.5 and 6.0 mg/mL protein identified three potential oligomers, first assigned as a hexamer, a nonamer and a large assembly beyond the resolution of the column. As shown in Figure 4, the relative distribution of these oligomers changes as the protein environment changes as a function of both pH and enzyme concentration. As the enzyme concentration increases from 0.7 to 6.0 mg/mL at pH 9.0 (Figure 4A), there is a clear increase in the peak height of the largest oligomer and a concurrent decrease in the peak height of the putative nonameric species, suggesting the protein favors the larger oligomer under higher protein concentrations, but some of the smaller putative nonamer peak remains present. The change in distribution between the available oligomeric states is more pronounced in response to change in pH (Figure 4B). At pH 7.5, each of three oligomers is present, with the putative nonameric species predominant, while the larger oligomer and the hexameric species are similar in abundance. As the pH increases to 8.3, the larger oligomer remains roughly at the same abundance, while the distribution of hexamer and putative nonamer shifts toward the hexamer. By contrast, at pH 9.0, there is almost entirely a single species, the larger oligomer, with a small amount of putative nonamer and no detectable hexamer remaining (Fig. 4B, red).


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Figure 4. Oligomeric state changes as a function of enzyme concentration and pH. (A) Size exclusion chromatography (SEC) performed at pH 9.0 with 6.0 mg/mL (red) and 0.7 mg/mL (blue) BcHMGR. Absorbance signals were normalized to the same scale despite the difference in enzyme concentrations in order to better compare relative peak heights. (B) Size exclusion chromatography performed at 6.0 mg/mL enzyme at pH 7.5 (blue), pH 8.3 (red) and pH 9.0 (green). From right to left, these peaks correspond to a hexamer, a potential nonamer, and a large particle beyond the size resolution of the SEC column. ANS Fluorescence Demonstrates a Ligand-Dependent Tertiary Structure Change The transition between quaternary forms of a morpheein proceeds through a dissociation event, followed by a conformational change and reassociation into a different quaternary form. The extrinsic probe 8-anilino-1-naphthalenesulfonic acid (ANS) demonstrates significantly increased fluorescence when exposed to hydrophobic environments and is commonly used as a solution probe for changes in structure as the protein environment changes (23). In order to characterize potential substrate-dependent structural changes in BcHMGR, NAD+ and coenzyme A were titrated into solutions of BcHMGR at 0.5, 1.0 and 2.0 mg/mL, pH 7.5, and ANS fluorescence monitored. Titration of BcHMGR with NAD+ or coenzyme A at varying concentrations of enzyme results in different degrees of ANS fluorescence and therefore structure change (Figure 5). At 0.5 mg/mL BcHMGR, both substrates result in an increase in ANS fluorescence, indicating some degree of increased access of the probe to hydrophobic regions of the protein. Coenzyme A results in a 55% increase in ANS fluorescence, while NAD+ results in a 30%


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Biochemistry

increase in ANS fluorescence (Figure 5A). This trend is reversed at 1.0 mg/mL BcHMGR (Figure 5B), and the magnitude of the effects is decreased (CoA: 17% increase; NAD+: 25% increase). As enzyme concentration increases to 2.0 mg/mL, however, the effects of NAD+ and coenzyme A are more like those at 0.5 mg/mL. At this enzyme concentration, coenzyme A induces a 40% increase, while NAD+ induces only a 15% increase (Figure 5C).

A

B

C

Figure 5. ANS fluorescence as a function of substrate concentration. (A) 0.5 mg/mL BcHMGR. (B) 1.0 mg/mL BcHMGR. (C) 2.0 mg/mL BcHMGR. In all plots, coenzyme A is represented in blue and NAD+ in green. Points represent the average of triplicate trials, with error bars representing one standard deviation.


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Crystal Structure of BcHMGR in Complex with Coenzyme A and ADP BcHMGR cocrystallized with coenzyme A crystallized in the P6322 space group with a monomer in the asymmetric unit. Upon structure determination, it was discovered that ADP was also present bound to the enzyme, though it was not added prior to or after crystallization, suggesting that it copurified with the enzyme. The BcHMGR monomer shares significant structural similarity with the monomeric units of other bacterial HMGRs whose crystal structures have been determined. Root-mean-square deviations (RMSDs) for C atoms range from 0.340 to 0.401 Å between Bc and PmHMGR (24, 25), from 0.337 to 0.457 Å between Bc and Delftia acidovorans (Da) HMGR (26), and from 0.772 to 0.893 Å between Bc and Streptococcus pneumoniae (Sp) HMGR (27). The stronger similarity between Bc, Pm, and DaHMGR is expected, as BcHMGR shares greater than 65% sequence identity with these two homologues, but only 41% with SpHMGR. Like these related structures, the BcHMGR monomer (Figure 6) is comprised of a small and large domain. The small domain corresponds to the non-Rossmann dinucleotide-binding domain in other bacterial HMGRs, which is a four-stranded antiparallel sheet with two crossover helices lining one side of the sheet. The large domain is mixed / and contains all the residues known to be required for catalysis (28).


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Biochemistry

Figure 6. Structure of the BcHMGR monomer bound to coenzyme A and ADP. Coenzyme A and ADP are rendered as sticks with carbons in yellow. The large domain binds coenzyme A and is positioned to the left of the figure. The small domain binds ADP and is positioned to the right of the figure. All structure images were produced with PyMOL.

Coenzyme A binds at the expected location, based on comparisons to other known bacterial HMGR structures, while ADP occupies the same binding site as the ADP moiety of NAD(P)H (25-27) in these structures. Like other bacterial HMG-CoA reductases, BcHMGR appears to have a disordered C-terminal domain, as the last 53 residues of the full-length protein could not reliably be placed into electron density. It is common for this domain to be disordered in bacterial HMGR structures, often requiring multiple substrates to be bound in order for the domain to order and close over the active site (25, 26).

Coenzyme A and ADP Binding in BcHMGR


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Coenzyme A binds to BcHMGR in the same location as it binds to other known bacterial HMGRs, bending around a large domain -helix with the pantetheine moiety extending toward the active site (Figure 7A). Indeed, the large domain provides all of the binding interactions between BcHMGR and coenzyme A. Arg11, highly conserved across HMGR sequences, forms a cation-pi interaction with the adenine ring of coenzyme A, as well as providing hydrogen bonding and electrostatic interactions with the bridging diphosphate. The Tyr67-OH and Lys95NH3 also provide hydrogen bonding and potential electrostatic (Lys95) binding interactions with the 3’-phosphate. These are the only direct protein-to-ligand contacts to coenzyme A, which suggests that binding is relatively weak, allowing for this substrate to associate and dissociate easily over the course of the reaction. Binding of ADP to BcHMGR is a surprise, as ADP was not added to the protein solution prior to or during crystallization. However, it binds at a familiar location, overlaying closely with the ADP moiety of NAD(H) in ternary complex structures of PmHMGR and the NAD+-bound structure of DaHMGR (25, 26). Unlike coenzyme A, ADP is bound entirely by the small domain. The adenine ring is coordinated by hydrogen bonds from both the backbone amide nitrogen and side chain of Asp183, while Thr189 hydrogen bonds with the 3’-OH of the ADP ribose. Other direct protein-to-ligand contacts are between the backbone amide nitrogens of Met185 and Gly186 and the ADP -phosphate. Asp146, which is conserved across other putative oxidative bacterial HMGRs including Da and PmHMGRs, also hydrogen bonds to the 3’-OH of the ADP ribose (Figure 7B). This residue has been implicated to help discriminate against NADP(H) in oxidative bacterial HMGRs (14) and forms a hydrogen bond with the 2’OH of the adenine ribose of NAD(H) in both Da and PmHMGR. It is possible that ADP and NAD(H) do not share this interaction due to the lack of the nicotinamide ring and its


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Biochemistry

accompanying ribose on ADP, leading to a slight change in positioning. Omit electron density maps for both substrates are shown in Figure S1.

Figure 7. Ligand binding sites in BcHMGR. Protein atoms are labeled with carbons in grey. ligand atoms. Coenzyme A (A) and ADP (B) are labeled as sticks with carbons in yellow. Dashed lines represent hydrogen bonding or salt bridge interactions. Oligomeric State of BcHMGR All known crystal structures of HMGR, regardless of species, are at least obligate dimers: a monomer is unable to form a complete active site, due to positioning of catalytically required residues in the monomer and binding sites for HMG-CoA/mevalonate and NAD(P)H. The monomeric structure of BcHMGR is also clearly unable to form a complete active site, with the catalytically important Asp283 over 17 Å away from the required proton donor/acceptor Lys267, a distance incompatible with its proposed catalytic role as a general acid/base (26, 28). In addition, at their closest approach, ADP and coenzyme A are over 34 Å away from one another on opposite sides of the protein. A 2-fold symmetry axis, however, demonstrates that BcHMGR also has a dimeric structure very similar to the dimers of other bacterial HMGRs (Figure 8A). Formation of the dimer buries 4605.1 Å of surface area as calculated by the PISA server (19), 54% of which comes from hydrophobic side chains. The interactions that comprise the dimer


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interface come primarily from large domain to large domain contacts with some large domain to small domain contacts, but the small domains do not contact one another. The 6-fold axis of the P6322 space group shows that BcHMGR also forms a trimer of dimers, or hexameric, state (Figure 8B), confirming one of the species identified in size exclusion chromatography experiments. This hexameric species is also seen in PmHMGR, which forms both dimers and hexamers in solution and in the crystal (24, 25, 29), and DaHMGR, which forms a similar trimer of dimers in the crystal (26). SpHMGR (27) and Listeria monocytogenes (Lm) HMGR (22) both appear to form no higher order form than dimers, as judged by crystal packing (SpHMGR) and size exclusion chromatography-multiangle light scattering (SEC-MALS) (LmHMGR). There is no crystallographic evidence of a nonamer in this structure, nor is there any immediately apparent trimeric species that could act as the lower-order intermediate between hexamer and nonamer in a potential morpheein transition. Formation of the hexamer buries a significantly smaller amount of surface area (991.3 Å2) and is more hydrophilic in nature, with only 38% of the interface contacts provided by hydrophobic side chains. The hexamer interface is formed between a crossover helix of one dimer resting across the four-stranded antiparallel -sheet of the large domain on an adjacent dimer, reminiscent of a cylinder resting in the palm of a hand (Figure 8C), with remarkably few interactions between neighboring dimers. Only a single salt bridge and two protein-protein hydrogen bonds hold the interface together. This interface is very similar to that seen in the two structures of putatively oxidative bacterial HMGRs (Pm and DaHMGR). Crystal packing analysis reveals a third interface, between the N-terminal regions of dimers involved in adjacent hexamers. This interface is not large (391.5Å, 41% hydrophobic) and is formed primarily by residues 8-17 in the crystal structure.


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Figure 8. Oligomeric states of BcHMGR present in the crystal. (A) A 2-fold symmetry axis generates a dimer. (B) The 63 screw symmetry axis generates a trimer of dimers or hexameric state as seen in size-exclusion chromatography. (C) A close up view of the hexamer interface shows the small domain helices of each dimer rest against the large domain antiparallel sheet of an adjacent dimer. Molecular Dynamics Simulations of N-Terminal Interface The N-terminal region involved in the hexamer-to-hexamer interface includes Arg11, which forms a cation-pi interaction with the coenzyme A adenine ring and a salt bridge with the bridging diphosphate (Figure 9A). Given that coenzyme A is able to induce double-saturation kinetics as demonstrated above and induces a change in hydrophobic accessibility to the ANS probe, this interface is of particular interest. To simulate the effects of removing coenzyme A from the crystal structure, molecular dynamics simulations were performed using the NAMD/VMD package. The simulation included two dimers involved in the interface of interest


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with coenzyme A manually edited out of the original coordinate file. These simulations revealed that in the absence of coenzyme A, Arg11 swings away from the coenzyme A-binding site and forms a bipartite salt bridge with Asp237 from the adjacent dimer (Figure 9B). This suggests that when coenzyme A is present, this salt bridge interaction is broken as Arg11 moves into binding position over the adenine ring and the interface is weakened, possibly promoting or allowing for the easier dissociation of the hexamer.


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Figure 9. Molecular dynamics simulation of N-terminal hexamer-to-hexamer interface. (A) The experimentally determined BcHMGR structure shows Arg11 closed over the coenzyme A adenine ring and interacting with the bridging diphosphate. (B) Molecular dynamics simulations in the absence of coenzyme A show Arg11 swinging away from the binding site to form a bipartite salt bridge with Asp237 from a protein chain in a neighboring hexamer. (C) Overlay of both structures, with color schemes consistent from (A) and (B).


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DISCUSSION The morpheein model of allostery describes a dynamic distribution of functionally distinct nonadditive oligomeric forms of a protein (5). In this model, changes in a wide range of factors including substrate concentration, pH, and protein concentration can result in a redistribution of these oligomeric states through a dissociation, tertiary structure change, and reassociation process. According to the morpheein model, each of the higher-order states can exhibit a different level of enzymatic activity, which allows for regulation of the enzyme in response to changes in physiological conditions. Earlier characterization of BcHMGR (13) indicated the enzyme possessed some of the characteristics typical of a morpheein. These included multiple oligomers at pH 7.5 as judged by size exclusion chromatography, each of which appeared to have different Vmax values, and double-saturation kinetics that could be explained by a substratedependent shift from a lower activity form to a higher activity form. We have now demonstrated that BcHMGR undergoes a number of structural and functional changes dependent upon substrate concentration, pH and enzyme concentration. Size exclusion chromatography demonstrates clearly that the distribution of oligomers changes as a function of pH and enzyme concentration. The transition from double-saturation kinetics at pH 7.5 and 8.3 to a more typical sigmoidal curve at pH 9.0 correlates with the single oligomer seen by size exclusion chromatography at that pH. Kinetic data at pH 9.0 suggests that catalysis is occurring from a single active state (Figure 2C) at a similar Vmax to pH 7.5 (Figure 2A), where the putative nonamer is more favored (Figure 3B), which might indicate that the large particle dissociates into nonamers in order to catalyze the oxidation of mevalonate. It is notable that at pH 9.0, decreasing the enzyme concentration results in a shift away from the largest oligomer toward the putative nonamer (Figure 4A), giving further support to the hypothesis that this large particle


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Biochemistry

forms from this potential nonameric species. Whether or not the large particle is physiologically relevant remains unknown, warranting further study as to whether it can form under milder conditions likely to be found inside a cell. Although pH 9.0 is far from physiological pH, it does mimic the effect that physiological signals may have on the protonation states of surface residues. The hexamer appears to be the most enzymatically active, as the highest Vmax is observed at pH 8.3 (Figure 2B), where the hexamer and putative nonamer are equally favored as judged by size exclusion chromatography (Figure 4B). Changes in any of the three substrates of the oxidative reaction (mevalonate, NAD+ and coenzyme A) in the saturating presence of the other two results in double-saturation kinetics at pH 7.5 (13). However, only two of these substrates exert any structural effect when binding singly to the enzyme. Given the relatively small change in ANS fluorescence upon addition of NAD+ to a solution of BcHMGR, it would appear that this substrate’s effects on the morpheein transition are primarily through changes in hydrophilic interfaces and the resulting quaternary structure change. Coenzyme A, on the other hand, induces a greater change in hydrophobic exposure, suggesting that it might exert more of an effect on tertiary structure. It is interesting to note that neither substrate has a marked effect on ANS fluorescence at 1.0 mg/mL BcHMGR, possibly suggesting the presence of a structural state resistant to the structural changes observed at other enzyme concentrations. Crystal structure and molecular dynamics evidence provides some insights into how substrates might affect a potential morpheein transition. By analogy to homologous structures, NAD+ would bind at the same site as ADP does in the structure described here (Figure S2). The small domain dinucleotide binding site is integral to the hexamer interface, and conformational changes involved in NAD+ binding could easily promote a modification of this interface. The PISA server suggests the hexamer interface in BcHMGR is


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not very stable, with a Gdiss of -1.1 kcal/mol, indicating that it is slightly thermodynamically favorable for this complex to dissociate. 62% of the contacting residues involved in this interface are hydrophilic, so if NAD+ is promoting the dissociation of this interface, we would not expect to see a major change in ANS fluorescence as a result. Coenzyme A binding, on the other hand, results in a more significant increase in hydrophobic exposure and ANS fluorescence. Molecular dynamics simulations do support the idea of coenzyme A resulting in a potential structural change. Binding of this substrate is predicted to induce the dissociation of a hexamer-to-hexamer interface via a salt bridge between the key binding residue Arg11 and Asp237 on the adjacent hexamer (Figure 9). This interface is relatively small (391.5 Å2) and primarily hydrophilic, making it unlikely that exposure of the two hexamers that make up the interface would account for the increase in ANS fluorescence. The stoichiometry of the putative nonamer remains uncertain. Despite the predicted instability of the hexamer and the fact that the putative nonamer is the more favored oligomer at pH 7.5, the hexamer is the species that crystallizes readily at this pH. Crystal packing may allow for a more stable hexamer, with interactions between neighboring hexamers providing additional thermodynamic stability to disfavor the dissociation event. In solution, it is plausible that the dissociation of the hexamer to a lower-order oligomer could provide an opportunity to undergo the necessary tertiary structure or conformational change required to promote a nonamer. However, the identity of this lower-order state also remains elusive. The smallest common factor of a hexamer and a nonamer is a trimer, but there is no current evidence for a trimeric species. It is not unusual for a morpheein to have a highly metastable lower-order transition state. Porphobilinogen synthase, the first identified morpheein, transitions between a highactivity octamer and a low-activity hexamer (30) through a dimeric intermediate only identified


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Biochemistry

initially through a mutation of the wild-type enzyme (31) due to its instability and later seen by size exclusion chromatography (32). The more significant puzzle, however, is how such a trimer of BcHMGR might form. Dissociation of the trimer-of-dimers hexamer seen in the crystal structure would likely result in a dimeric structure, and all available HMGR structures strongly suggest that the minimum active unit of the enzyme is a dimer. It is conceivable that the nonamer could instead be an octamer or decamer. The original molecular weight determination of this size exclusion peak corresponded to 9.3±0.2 monomers (13) of BcHMGR. The inherent uncertainty of molecular weight determination by comparison to standards does suggest that this putative nonamer may indeed be a decamer or even a dodecamer, given the likelihood that it would form from a dimeric intermediate. Crystallization of the apoenzyme at pH 7.5, where this putative nonamer is more favored, may provide further clues as to the identity of this species. Other, higher resolution techniques such as SEC-MALS, analytical ultracentrifugation, or smallangle x-ray scattering should also provide further evidence regarding the identity of the other oligomers observed by SEC alone. From kinetics data and tertiary and quaternary structure change evidence, it is increasingly likely that BcHMGR functions as a morpheein. What forces are at work to drive this potential morpheein behavior in BcHMGR, not observed in the closely related (72% sequence identity) PmHMGR? There are clear differences between the activity and structure of these two enzymes: PmHMGR exhibits hyperbolic kinetics in all published cases, with no evidence of a regulatory scheme, and is in equilibrium between a dimer and a hexamer in solution. One possible explanation may come from the stability of the hexameric assembly. While the BcHMGR hexamer is predicted to be slightly thermodynamically unstable as mentioned above, the PmHMGR apoenzyme (PDB ID 4I64) structure (33) is predicted to form a much more stable


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hexamer (Gdiss = 10.1 kcal/mol). Likewise, DaHMGR has an even more stable hexameric assembly (Gdiss = 15.6 kcal/mol). For both Pm and DaHMGR, the PISA server predicts the hexamer to be the most stable assembly, while for BcHMGR, it predicts the dimer to be the most stable assembly. Given how important dissociation of the higher-order forms is for the morpheein transition, this metastable hexameric assembly could provide BcHMGR an opportunity for structural rearrangement that other related HMGRs may not have. A detailed examination of the hexamer interfaces of Bc, Pm, and DaHMGR suggests that BcHMGR has significantly fewer direct protein-protein contacts across the interface: only one salt bridge (Arg128 to Glu320) and two additional hydrogen bonding interactions. By comparison, PmHMGR also has the Arg128-Glu320 salt bridge, but seven additional hydrogen bonding interactions. DaHMGR, which has the strongest interface, has two potential salt bridges (Arg128-Glu320 and Arg226-Asp123) and eight additional hydrogen bonds. DaHMGR is the only one of the three with an arginine at position 226 (BcHMGR has an aspartate, while PmHMGR has a glutamine), which makes it the only one able to form the second salt bridge (Figure 10, left). In addition, the hexamer interface in BcHMGR is slightly more hydrated (five water molecules) than either Pm or DaHMGR (three each), as would befit an assembly that dissociates more readily (Figure 10, right). It is interesting to note that PBGS also demonstrates similar phylogenetic diversity, in which some species (humans, for example) express a morpheein form of the enzyme, while others do not, despite significant sequence identity (>35%) (34).


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Biochemistry

Figure 10: Comparison of Hexamer Interfaces. The hexamer interfaces of BcHMGR (greens), PmHMGR (blues), and DaHMGR (yellows) are shown, aligned on the basis of the small domain helix. (A) All three structures share the Arg128-Glu320 salt bridge, but only DaHMGR can form the Arg226-Asp123 salt bridge. (B) BcHMGR has more bound water molecules at the interface between the small domain helix and the large domain sheet at the hexamer interface. The very large oligomer appears to be unique to BcHMGR. Based on the molecular dynamics simulations described here, it is possible that substrate binding may drive the association and dissociation of this large species. Only BcHMGR has an aspartate residue (Asp237) at the proper position for Arg11 to form a salt bridge interaction between neighboring hexamers in the absence of coenzyme A (Figure 9). Indeed, among the five HMGRs whose oligomeric states are known or inferred from crystal packing, not only is Asp237 not conserved, only BcHMGR has a negatively charged residue at this position. Taken together, the data suggest an initial map for a potential morpheein transition in BcHMGR. The hexamer appears to be the most active state, as judged by pH-dependent kinetics and size exclusion chromatography data. Conversely, conditions that tend to favor the putative nonamer (high pH and high enzyme concentrations) promote a relatively lower activity. NAD+ appears to


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be more important for driving the dissociation of higher-order oligomers toward the lower-order transition state, as judged by relatively minor changes in hydrophobic exposure and the nature of the hexameric interface seen in the crystal. Coenzyme A has a stronger effect on hydrophobic surface exposure and may be primarily responsible for driving the transition between states that determine the eventual assembly into the higher-order active hexamer and putative nonamer. The largest oligomer appears to assemble, at the very least, from the putative nonameric species as judged by size exclusion chromatography as a function of enzyme concentration. Interfaces present between hexamers in the crystalline state and corresponding molecular dynamics simulations suggest that the large oligomer might also form from the hexameric species. In summary, we have now further characterized a number of factors that modulate a potential morpheein transition between active quaternary forms of BcHMGR. Kinetic assays demonstrate a transition to single state kinetics as the pH of the solution increases, and a decrease in kcat as the enzyme concentration increases. We have also demonstrated that BcHMGR undergoes substrate-dependent structure change as judged by ANS fluorescence, and dramatic changes in the distribution of quaternary forms in response to both pH and enzyme concentration. The crystal structure of the hexamer in complex with coenzyme A and ADP bound at the NAD(H) site, combined with molecular dynamics simulations, provides a possible mechanism by which substrate-dependent shifts in quaternary structure may occur, and suggests why BcHMGR appears to be unique among other characterized HMG-CoA reductases in possessing such complex behavior. Classification of BcHMGR as a morpheein will require demonstration of similar changes as a function of substrate concentration under physiological conditions, but the demonstration of dynamic changes in oligomeric state and activity provided here correlate with the behavior expected for a morpheein. Successful classification of Burkholderia cenocepacia


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HMG-CoA reductase as a morpheein would add a new member to this fascinating family of structurally and functionally complex enzymes.


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AUTHOR INFORMATION Corresponding Author * E-mail: [email protected] ORCID Jeffrey Watson: 0000-0002-6939-0137 Present Addresses 1 Current

address: Department of Chemistry and Biochemistry, University of California, San

Diego, San Diego, CA, 92093, USA 2

Current address: Department of Biophysics and Biophysical Chemistry, Johns Hopkins

University, Baltimore, MD, 21287, USA Author Contributions J.M.W. conceived of the project and designed the experiments together with R.B.P., C.W.H., S.M.D. and S.W.A.S. J-C.A. performed molecular dynamics experiments. A.M.W. assisted with crystallization and collected x-ray data. A.M.W. and K.M.L. determined and refined the crystal structure. J.M.W., R.B.P., C.W.H., S.M.D., S.W.A.S. and J-C.A. analyzed the data. J.M.W. wrote the paper. Funding Sources This research was supported by the Gonzaga Science Research Program and a grant to Gonzaga University from the Howard Hughes Medical Institute through the Undergraduate Science Education Program.


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Notes The authors declare no competing financial interest. ABBREVIATIONS HMG-CoA, 3-hydroxy-3-methylglutaryl coenzyme A; HMGR, HMG-CoA reductase; ANS, 8anilino-1-naphthalenesulfonic acid; RMSD, root-mean-square deviations; PDB, Protein Data Bank. ACCESSION CODES Burkholderia cenocepacia HMG-CoA reductase: UniProt KB B4EKH5

SUPPORTING INFORMATION Omit maps of coenzyme A and ADP bound to BcHMGR. Overlay of ADP and NAD+ binding in bacterial HMGRs.

REFERENCES References 1. Changeux, J. P. Allostery and the Monod-Wyman-Changeux model after 50 years. Annu. Rev. Biophys. (2012), 41, 103-133. 2. Koshland, D. E.; Nemethy, G.; Filmer, D. Comparison of experimental binding data and theoretical models in proteins containing subunits . Biochemistry (1966), 5, 365-385. 3. Brunori, M. Variations on the theme: allosteric control in hemoglobin. FEBS J. (2014), 281, 633-643. 4. Zancan, P.; Marinho-Carvalho, M. M.; Faber-Barata, J.; Dellias, J. M.; Sola-Penna, M. ATP and fructose-2,6-bisphosphate regulate skeletal muscle 6-phosphofructo-1-kinase by altering its quaternary structure. IUBMB Life (2008), 60, 526-533.


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5. Jaffe, E. K. Morpheeins--a new structural paradigm for allosteric regulation. Trends Biochem. Sci. (2005), 30, 490-497. 6. Jaffe, E. K. The Remarkable Character of Porphobilinogen Synthase. Acc. Chem. Res. (2016), 49, 2509-2517. 7. Jaffe, E. K.; Lawrence, S. H. Allostery and the dynamic oligomerization of porphobilinogen synthase. Arch. Biochem. Biophys. (2012), 519, 144-153. 8. Selwood, T.; Jaffe, E. K. Dynamic dissociating homo-oligomers and the control of protein function. Arch. Biochem. Biophys. (2012), 519, 131-143. 9. Mouilleron, S.; Badet-Denisot, M. A.; Pecqueur, L.; Madiona, K.; Assrir, N.; Badet, B.; Golinelli-Pimpaneau, B. Structural basis for morpheein-type allosteric regulation of Escherichia coli glucosamine-6-phosphate synthase: equilibrium between inactive hexamer and active dimer. J. Biol. Chem. (2012), 287, 34533-34546. 10. Jaffe, E. K.; Stith, L.; Lawrence, S. H.; Andrake, M.; Dunbrack, R. L. A new model for allosteric regulation of phenylalanine hydroxylase: implications for disease and therapeutics. Arch. Biochem. Biophys. (2013), 530, 73-82. 11. Korasick, D. A.; Gamage, T. T.; Christgen, S.; Stiers, K. M.; Beamer, L. J.; Henzl, M. T.; Becker, D. F.; Tanner, J. J. Structure and characterization of a class 3B proline utilization A: Ligand-induced dimerization and importance of the C-terminal domain for catalysis. J. Biol. Chem. (2017), 292, 9652-9665. 12. Snasel, J.; Pichova, I. Allosteric regulation of pyruvate kinase from Mycobacterium tuberculosis by metabolites. Biochim. Biophys. Acta Proteins Proteom (2019), 1867, 125139. 13. Schwarz, B. H.; Driver, J.; Peacock, R. B.; Dembinski, H. E.; Corson, M. H.; Gordon, S. S.; Watson, J. M. Kinetic characterization of an oxidative, cooperative HMG-CoA reductase from Burkholderia cenocepacia. Biochim. Biophys. Acta (2014), 1844, 457-464. 14. Friesen, J. A.; Lawrence, C. M.; Stauffacher, C. V.; Rodwell, V. W. Structural determinants of nucleotide coenzyme specificity in the distinctive dinucleotide binding fold of HMGCoA reductase from Pseudomonas mevalonii. Biochemistry (1996), 35, 11945-11950. 15. Otwinowski, Z.; Minor, W. Processing of X-ray diffraction data collected in oscillation mode. Methods Enzymol. (1997), 276, 307-326. 16. Adams, P. D.; Afonine, P. V.; Bunkoczi, G.; Chen, V. B.; Davis, I. W.; Echols, N.; Headd, J. J.; Hung, L. W.; Kapral, G. J.; Grosse-Kunstleve, R. W.; McCoy, A. J.; Moriarty, N. W.; Oeffner, R.; Read, R. J.; Richardson, D. C.; Richardson, J. S.; Terwilliger, T. C.; Zwart, P. H. PHENIX: a comprehensive Python-based system for macromolecular structure solution. Acta Crystallogr. D Biol. Crystallogr. (2010), 66, 213-221.


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17. Emsley, P.; Cowtan, K. Coot: model-building tools for molecular graphics. Acta Crystallogr. D Biol. Crystallogr. (2004), 60, 2126-2132. 18. Phillips, J. C.; Braun, R.; Wang, W.; Gumbart, J.; Tajkhorshid, E.; Villa, E.; Chipot, C.; Skeel, R. D.; Kale, L.; Schulten, K. Scalable Molecular Dynamics with NAMD . Journal of Computational Chemistry (2005), 26, 1781-1802. 19. Krissinel, E.; Henrick, K. Inference of macromolecular assemblies from crystalline state. J. Mol. Biol. (2007), 372, 774-797. 20. Best, R. B.; Zhu, X.; Shim, J.; Lopes, P. E.; Mittal, J.; Feig, M.; Mackerell, A. D. Optimization of the additive CHARMM all-atom protein force field targeting improved sampling of the backbone phi, psi and side-chain chi(1) and chi(2) dihedral angles. J. Chem. Theory Comput. (2012), 8, 3257-3273. 21. Jorgensen, W. L.; Chandrasekar, J.; Madura, J. D.; Impey, R. W.; Klein, M. L. Comparison of simple potential functions for simulating liquid water . Journal of Chemical Physics (1983), 79, 926-935. 22. Theivagt, A. E.; Amanti, E. N.; Beresford, N. J.; Tabernero, L.; Friesen, J. A. Characterization of an HMG-CoA reductase from Listeria monocytogenes that exhibits dual coenzyme specificity. Biochemistry (2006), 45, 14397-14406. 23. Hawe, A.; Sutter, M.; Jiskoot, W. Extrinsic fluorescent dyes as tools for protein characterization. Pharm. Res. (2008), 25, 1487-1499. 24. Lawrence, C. M.; Rodwell, V. W.; Stauffacher, C. V. Crystal structure of Pseudomonas mevalonii HMG-CoA reductase at 3.0 angstrom resolution. Science (1995), 268, 1758-1762. 25. Tabernero, L.; Bochar, D. A.; Rodwell, V. W.; Stauffacher, C. V. Substrate-induced closure of the flap domain in the ternary complex structures provides insights into the mechanism of catalysis by 3-hydroxy-3-methylglutaryl-CoA reductase. Proc. Natl. Acad. Sci. U. S. A. (1999), 96, 7167-7171. 26. Ragwan, E. R.; Arai, E.; Kung, Y. New Crystallographic Snapshots of Large Domain Movements in Bacterial 3-Hydroxy-3-methylglutaryl Coenzyme A Reductase. Biochemistry (2018), 57, 5715-5725. 27. Miller, B. R.; Kung, Y. Structural Features and Domain Movements Controlling Substrate Binding and Cofactor Specificity in Class II HMG-CoA Reductase. Biochemistry (2018), 57, 654-662. 28. Hedl, M.; Tabernero, L.; Stauffacher, C. V.; Rodwell, V. W. Class II 3-hydroxy-3methylglutaryl coenzyme A reductases. J. Bacteriol. (2004), 186, 1927-1932.


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29. Rogers, K. S.; Rodwell, V. W.; Geiger, P. Active form of Pseudomonas mevalonii 3hydroxy-3-methylglutaryl coenzyme A reductase. Biochem. Mol. Med. (1997), 61, 114-120. 30. Breinig, S.; Kervinen, J.; Stith, L.; Wasson, A. S.; Fairman, R.; Wlodawer, A.; Zdanov, A.; Jaffe, E. K. Control of tetrapyrrole biosynthesis by alternate quaternary forms of porphobilinogen synthase. Nat. Struct. Biol. (2003), 10, 757-763. 31. Tang, L.; Breinig, S.; Stith, L.; Mischel, A.; Tannir, J.; Kokona, B.; Fairman, R.; Jaffe, E. K. Single amino acid mutations alter the distribution of human porphobilinogen synthase quaternary structure isoforms (morpheeins). J. Biol. Chem. (2006), 281, 6682-6690. 32. Selwood, T.; Tang, L.; Lawrence, S. H.; Anokhina, Y.; Jaffe, E. K. Kinetics and thermodynamics of the interchange of the morpheein forms of human porphobilinogen synthase. Biochemistry (2008), 47, 3245-3257. 33. Steussy, C. N.; Critchelow, C. J.; Schmidt, T.; Min, J. K.; Wrensford, L. V.; Burgner, J. W.; Rodwell, V. W.; Stauffacher, C. V. A novel role for coenzyme A during hydride transfer in 3-hydroxy-3-methylglutaryl-coenzyme A reductase. Biochemistry (2013), 52, 5195-5205. 34. Jaffe, E. K. The remarkable character of porphobilinogen synthase. Acc. Chem. Res. (2016), 49, 2509-2517.


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Structural and functional characterization of dynamic

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