Protein Scaffold Activates Catalytic CO2 Hydrogenation by a Rhodium

Research Article Cite This: ACS Catal. 2019, 9, 620−625

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Protein Scaffold Activates Catalytic CO2 Hydrogenation by a Rhodium Bis(diphosphine) Complex Joseph A. Laureanti,† Garry W. Buchko,†,‡ Sriram Katipamula,† Qiwen Su,† John C. Linehan,† Oleg A. Zadvornyy,§ John W. Peters,†,§ and Molly O’Hagan*,†,⊥ †

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Fundamental and Computational Sciences Directorate, Pacific Northwest National Laboratory, Richland, Washington 99352, United States ‡ School of Molecular Biosciences, Washington State University, Pullman, Washington 99164, United States § Institute of Biological Chemistry, Washington State University, Pullman, Washington 99164, United States S Supporting Information *

ABSTRACT: The utilization of CO2 to generate chemical fuels, such as formic acid, is a potentially beneficial route to balance carbon emissions and reduce dependence on fossil fuels. The development of efficient catalysts for CO2 hydrogenation is needed to implement this fuel generation. In the molecular catalyst design presented here, we covalently attached a rhodium complex, ([RhI(PNglyP)2]−, where − PNglyP is defined as PEt2−CH2−N(CH2CO2 )−CH2−PEt2) to a protein scaffold, (lactococcal multidrug resistant regulator from Lactococcus lactis) to use the protein environment around the metal center to control substrate delivery and therefore enable and improve catalytic activity. The reactivities of the rhodium complex and the synthetic metalloenzyme were characterized by high-pressure operando NMR techniques. In solution, the rhodium complex alone is not a catalyst for CO2 hydrogenation. Incorporation of the rhodium complex into the protein scaffold resulted in a gain of function, turning on CO2 hydrogenation activity. The metalloenzyme displayed a turnover frequency of 0.38 ± 0.03 h−1 at 58 atm and 298 K and achieved an average turnover number of 14 ± 3. Proposed catalytic intermediates generated and characterized suggest that the protein scaffold enables catalysis by facilitating the interaction between CO2 and the hydride donor intermediate. KEYWORDS: protein scaffold, carbon dioxide hydrogenation, rhodium complex, metalloenzyme

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intricate cooperativity between all coordination spheres of the catalytic center as well as the medium in this molecular system mirrors that of natural enzymes where the dynamic protein scaffold facilitates efficient reactivity at a metalloenzyme active site through interactions well beyond the metal center. In this study, instead of intricate ligand and medium design to achieve efficient substrate delivery, we utilize covalent attachment of a synthetic complex to a genetically encodable protein scaffold. This approach allows for bioengineering of the scaffold to identify the effects of outer coordination sphere interactions on reactivity.12,13 Specifically, we investigate the ability of a protein scaffold to modulate the interaction between CO2 and a catalytically inactive rhodium bis(diphosphine) complex, ([RhI(PNglyP)2]− where PNglyP is defined as PEt2−CH2−

ydrogenation of CO2 to formate using H2 generated from renewable sources is an attractive transformation for fuel production due to the high energy density of formic acid.1,2 The development of efficient catalysts for this transformation is required for the widespread generation of this fuel. The most active synthetic molecular catalysts for this transformation typically require noble metals and elevated temperatures and pressures.3−5 Contrary to this, the metal-dependent formate dehydrogenase enzymes (FDHs) readily catalyze the reduction of CO2 to formate at ambient temperatures and pressures using earth-abundant metals at their active sites.6 The outer coordination sphere of the metal-dependent FDH active site is proposed to control substrate delivery by hydrogen bonding interactions stabilizing the interaction between the hydride donor and the CO2 substrate during catalytic turnover.7 In this study, we look to emulate this interaction in a synthetic system by engineering a protein scaffold to control substrate delivery and thereby improve catalytic performance. Controlled substrate delivery has been shown to dramatically enhance the performance of synthetic catalysts.8,9 Recently, cooperativity between the primary, secondary, and outer coordination spheres as well as the medium has been used to mediate the rates of proton delivery within nickel catalysts for H2 production to achieve rate enhancements up to four orders of magnitude compared to first-generation catalyst design.10,11 The © XXXX American Chemical Society

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N(CH2CO2 )−CH2−PEt2). Incorporation of this catalytically incompetent rhodium complex into the protein scaffold results in a gain of function where the conjugated system is an artificial metalloenzyme for CO2 hydrogenation. The [RhI(PNglyP)2]− complex, 1 (Scheme 1), was chosen for study because the hydride donor ability of the Received: July 5, 2018 Revised: November 12, 2018

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DOI: 10.1021/acscatal.8b02615 ACS Catal. 2019, 9, 620−625

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ACS Catalysis

Scheme 1. Cartoon Representation for the Strategy to Covalently Attach 1 at LmrR* to Form the Conjugated Complex 2a

a

Equivalents refers to moles of Rh compared to moles of LmrR* as a dimer.

[RhIII(H)2(PNglyP)2]− was estimated to be poised to hydrogenate CO2 in water at pH 8.15 using previously published metrics, with details of the estimations outlined in the Supporting Information.14,15 The ligand and metal complex were synthesized as described in the Supporting Information, with procedures similar to those reported for related complexes.16,17 The lactococcal multidrug resistant regulator from Lactococcus lactis (LmrR) was chosen as the protein scaffold because it has a solvent-accessible cavity shown to accommodate various organic molecules and inorganic complexes.18−22 The LmrR sequence was modified with two mutations in the DNA binding domain to facilitate purification (K55D and K59Q) and M89C substitution.18 This variant is denoted as LmrR*. Expression and purification procedures for LmrR* are detailed in the Supporting Information. The maleimide linkage strategy was used to covalently attach 1 to C89 in LmrR*, as illustrated in Scheme 1. Cysteine-89 was chosen for the attachment point because its location would position the complex within the subunit cavity and has been shown not to disrupt the protein fold and dimeric assembly.23 For conjugation reactions, in situ peptide coupling chemistry was used to install the N-(2-aminoethyl)maleimide linkage onto the glycine residues of the [RhI(PNglyP)2]− complex. After amide bond formation, the solution was added to a dilute solution of LmrR*. The conjugated protein solution was then concentrated and unbound complex 1 was removed using a desalting NAP-5 column (GE Healthcare). Detailed reaction conditions can be found in the Supporting Information. Covalent attachment of the rhodium complex to LmrR* to form the metalloprotein, 2, was characterized using UV/visible and circular dichroism spectroscopy, electrospray ionization mass spectrometry (ESI-MS), inductively coupled plasma optical emission spectrometry (ICP-OES), and multidimensional nuclear magnetic resonance techniques. LmrR* is a homodimer that provides a single cysteine attachment point per monomer. Each PNglyP ligand has one maleimide; therefore, there is a possibility of 2:2 or 2:1 monomer:rhodium complex stoichiometries. In the 2:2 stoichiometry, each C89 is coupled to a different rhodium complex. In the 2:1 stoichiometry, a single rhodium complex can span the pocket attaching at C89 of each monomer. Complex 1 has characteristic charge transfer absorption at 389 nm with a measured extinction coefficient of 2250 M−1 cm−1. The coupling efficiency to LmrR* was determined using this unique 389 nm UV absorption band (Figure 1) to determine the metal content in the product. The protein content was determined using the corrected 280 nm UV absorption band and verified with a Bradford assay. The reaction conditions could be controlled to generate a product (2) with a coupling efficiency in the 96% range. The high coupling efficiency relative

Figure 1. UV−vis absorbance spectra of 1 (black), 2 (red), apo-LmrR* before the coupling reaction (gray), and purified LmrR* post exposure to 1 and peptide coupling reagents (blue). All spectra were collected under 1.0 atm of N2.

to the monomer dictates that the vast majority of the metalloprotein product has a 2:2 monomer:rhodium complex stoichiometry. The use of excess amounts of 1 in the coupling reaction did not affect the coupling efficiency over 96%, suggesting that there is no adventitious rhodium binding and the desalting procedure is efficient at removing any complex that is not covalently bound to the protein. Additionally, exposure of a mixture of 1 and peptide coupling reagents (without maleimide) to LmrR* followed by identical reaction and workup conditions as above results in a colorless product and a UV/vis spectrum lacking the 389 nm band (blue spectrum, Figure 1). Inductively coupled plasma optical emission spectrometry found 0.96 ppm rhodium in a sample of 2 prepared at 1.0 ppm rhodium based on the extinction coefficient. The S:Rh ratio was 4.8:1, near the 4:1 expected ratio in a complex with 2:2 monomer:rhodium stoichiometry. Also investigated was the P:Rh ratio, and it was found to be 4.2:1, near the 4:1 ratio expected. The most abundant peak in the ESIMS data was an LmrR* monomer with the maleimide attached PNglyP ligand without rhodium: observed 14022.62 m/z, calculated 14020.81 m/z, abundance 100%. The LmrR* monomer with a complete covalently attached rhodium complex is only 1.45% abundant: observed 14525.72 m/z, calculated 14525.14 m/z. The low abundance of the complete rhodium complex bound to LmrR* is attributed to its oxygen sensitivity where rhodium is lost during exposure to oxygen and further ionization in air.24 Covalent attachment of 1 to the protein scaffold did not significantly perturb the secondary structure, as illustrated by the nearly identical circular dichroism spectra before and after coupling, Figure S1A. The stability of the protein was altered upon attachment of 1 as the irreversible thermal denaturation curve shifts from TM ≈ 341 K in the apo protein to TM ≈ 327 K in 2, Figure S1B. Variable temperature 621

DOI: 10.1021/acscatal.8b02615 ACS Catal. 2019, 9, 620−625

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1 to the protein scaffold results in a gain of function, creating an active artificial metalloenzyme for CO2 hydrogenation. The catalytic activity of 2 under various conditions is summarized in Table 1. The turnover frequency (TOF) was calculated based on

circular dichroism did not show any significant contribution from Rh between 283 to 333 K, Figure S2. NMR spectroscopy was used to characterize the structure and reactivity of 1 and 2. The 1H−15N HSQC spectra of fully 15Nlabeled LmrR* and 2 are compared in Figure S3. The majority of the resonances are unperturbed after attachment of 1, providing additional evidence that coupling did not alter the protein’s overall structure significantly. However, as labeled in Figure S3, five pairs of amide cross peaks shift significantly (blue circles) and six new ones appear (cyan circles), indicating local perturbations to the protein’s structure, as might be expected with the covalent addition of 1. Product 2 exhibits a single resonance in the 31P{1H} spectrum, 2.35 ppm, with a very similar chemical shift as the resonances observed for 1, 2.20 ppm, under identical conditions, Figure 2. The 31P{1H}

Table 1. Reactivity Summary of 1 and 2a complex 1 1 2 2 2 2 2 2 2 1 + LmrR*c

pressure (atm) temperature (K) CO2:H2 34 58 1.7 17 34 58 34 34 17 58

298 298 298 298 298 298 323 353 298 298

1:1 1:1 1:1 1:1 1:1 1:1 1:1 1:1 0:1 1:1

rate (h−1)b nr trace 0.02 0.14 ± 0.02 0.23 ± 0.04 0.38 ± 0.03 0.08 nr nr nr

a

Conditions were 0.3 mM complex and 0.5 M sodium bicarbonate at pH 8.15 with sodium propane sulfonate as an internal integration standard. bRates were determined at a time point of 16 h. nr indicates no detectable product formation. c1, peptide coupling reagents, LmrR* without maleimide for covalent attachment.

the concentration of rhodium determined by our calculated extinction coefficient. As shown in the example in Figure 3A, the

Figure 2. 31P{1H} NMR spectra at room temperature in 0.5 M sodium bicarbonate pH 8.15 of (A) 1, (B) 2, (C) 1 under a H2 atmosphere, and (D) 2 under a H2 atmosphere.

spectrum for 2 is much broader than that for 1 due to the association of the complex to a ∼28 kDa protein dimer. In addition to further verifying successful coupling, Figure 2 suggests that the rhodium coordination environment does not change significantly upon attachment to the scaffold. Efforts to obtain a diffraction quality crystal for the 2:2 complex (2) using multiple screening conditions under anoxic conditions were unsuccessful. However, by adjusting the coupling conditions in Scheme 1, it was possible to predominately generate a product with a 2:1 monomer:rhodium complex stoichiometry. Diffraction quality crystals for the 2:1 product were obtained using the hanging drop technique, as detailed in the Supporting Information. The crystal structure resolved to 2.8 Å and showed the rhodium complex residing within the pocket of the LmrR* dimer with little perturbation to the overall protein tertiary structure relative to the native protein, Figure S4.17 Disorder is observed for the ligand and maleimide linkage, but the distance between the resolvable amine nitrogen of the PNglyP ligand and the thiol of C89 is sufficient to accommodate the maleimide linkage on both sides, Figure S5. High-pressure operando 1H NMR experiments using PEEK NMR cells25,27 were used to determine the reactivity of 1 and 2 under various conditions. Importantly, compound 1 is not a catalyst for CO2 hydrogenation under the conditions tested. No formate was produced, that could be detected, by 1 at 34 atm, and only trace amounts were produced at 58 atm. Only at 58 atm over the course of seven days was a substoichiometric formation of formate, 0.2 equiv, observed. Contrary to this, conjugation of

Figure 3. (A) Representative 1H NMR spectra showing the increase in the formate resonance over time for the reaction performed under 58 atm. (B) Plot of the rate of formate production (measured at 16 h) versus the pressure of CO2:H2. The inset is a plot of a representative experiment with data collected at 34 atm and also includes error bars as the standard deviation of three independent trials.

catalytic CO2 hydrogenation was quantitated by the appearance and growth of the formate resonance centered at 8.45 ppm in the 1 H NMR spectra. A plot of this representative data, shown in the inset in Figure 3B, indicates that the catalytic activity begins to level with time, perhaps due to protein oxidation as suggested by the buildup of covalent protein dimers over the course of the reaction (Figure S6). The metalloenzyme 2 exhibits a TOF of 0.38 h−1 at 58 atm and 298 K and achieved an average turnover number of 14 ± 3. No formate formation is observed for LmrR* exposed to a mixture of 1 and peptide coupling reagents (without maleimide), discussed above. As shown in Figure 3B, the catalytic rate of 2 exhibits a firstorder dependence on total CO2:H2 pressure, and as shown in Table 1, no formate formation is observed in the absence of added CO2 pressure. Catalytic activity was monitored at 1.7, 17, 34, and 58 atm, and representative data for catalytic runs are 622

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possible in water. Even at pH 11, well above the pH of catalytic experiments, [RhIII(H)2(PNglyP)2]− could not be deprotonated, Figure S9. This reactivity suggested that [RhIII(H)2(PNglyP)2]− could be the hydride donor. The [RhIII(H)2(PNglyP)2]− as the hydride donor is more similar to the previously proposed mechanism for [(H2)CoIII(dmpe)2]+ in aqueous media where cobalt(III)-dihydride is the hydride donor in aqueous media rather than cobalt(I)-monohydride as observed in organic solvents.15 The hydricity estimate of [RhIII(H)2(PNglyP)2]− is 32.2 kcal/ mol, which is above the 25.5 kcal/mol required for thermodynamically favoring CO2 hydrogenation. The estimated hydricity value is consistent with the observation that complex 1 is not a competent catalyst for CO2 hydrogenation. However, the conjugated protein, 2, is a competent catalyst for CO2 hydrogenation. This result suggests two possibilities: (1) the protein scaffold alters the hydricity of the RhIII-dihydride intermediate or (2) it changes the identity of the hydride donor to the RhI-monohydride. Estimating the hydricity of the RhImonohydride yields 12.2 kcal/mol, which positions this species to be a favorable hydride donor to CO2. Unfortunately, the deprotonation event required to form the RhI-monohydride is not possible in water under catalytic conditions as discussed above. The protein scaffold of 2 could alter the pKa of the RhIIIdihydride intermediate. However, the remarkably large change in pKa (>8 pKa units) required seems unlikely especially due to the solvent-exposed cavity in which the complex is attached. The RhIII-monohydride species is generated independently by protonation of 1 but is only observable at low pH, as characterized by 1H and 31P NMR spectroscopy, Figure 4. At

shown in Figure S7. Carbon-13-labeled carbon dioxide results in production of 13C-labeled formate (resonance at 171.8 ppm) under 1.7 atm of 1:1 13CO2:H2, Figure S8. While these results suggest that dissolved CO2 is the hydrogenation substrate, we cannot be definitive as CO2 and bicarbonate are in equilibrium under the conditions used. We have observed that in the absence of bicarbonate no catalysis is observed. This is most likely due to the need for a higher solution pH (neutral to basic) than can be achieved when pressurizing with CO2 (acidic). Using other buffers such as phosphate and tris (data not shown) at similar pH values, no formate is observed. It has previously been observed that nickel phosphines do not catalyze CO2 hydrogenation in the presence of phosphate buffer but do in the presence of bicarbonate buffer.26 A proposed catalytic cycle for CO2 hydrogenation by 2 is shown in Scheme 2. Upon addition of H2, the RhI square-planar Scheme 2. Proposed Catalytic Cycle of CO2 Hydrogenation at the Active Site of 2

complex forms an octahedral RhIII-cis-dihydride, which acts as the hydride donor to the substrate, CO2, to form a RhIIImonohydride species. The monohydride rhodium(III) complex is then deprotonated by a base in solution to yield the starting square-planar rhodium(I) complex. The RhI square-planar species is observed when either 1 or 2 is dissolved in bicarbonate buffered aqueous solutions at pH 8.15 by 31P{1H} NMR spectroscopy, Figure 2A,B. Adding H2 pressure then produces the cis-rhodium-dihydride, Figure 2C,D. Phosphorus-31 resonances are observed that are consistent with formation of the RhIII-cis-dihydride complex. As with the RhI intermediate, the resonances observed for the RhIII-dihydride species are very similar in chemical shift for both 1, 14.94 and −7.14 ppm, and 2, 15.97 and −6.01 ppm; Figure 2C,D, respectively. This proposed scheme is different than the mechanism previously proposed for rhodium analogues17,28 in organic media as well as the cobalt analogue, [(H2)CoIII(dmpe)2]+, in aqueous media.15 The hydride donor intermediate proposed for similar rhodium catalysts in organic media is a RhI-monohydride formed after deprotonation of the RhIII-dihydride. The reported pKa values of similar RhIII-dihydrides are estimated to be very high because they require very strong bases such as Verkade’s base, pKa conjugate acid = 33, for catalysis to occur.17,28 On the basis of these data, the pKa of 1 was estimated to be >16 in water. Consistent with this pK a , deprotonation of the [RhIII(H)2(PNglyP)2]− to form the RhI-monohydride was not

Figure 4. pH titrations of 1 to form the monohydride species monitored by (A) 31P{1H} and (B) 1H NMR spectroscopy. Conditions: 0.1 M MesHEPES buffer starting at pH 3.5, r.t.

pH < 6.5, a new hydride resonance at −21.8 ppm with JRh−H and JP−H of 15 Hz in the 1H NMR spectra is observed as well as its corresponding resonance in the 31P NMR spectrum with JRh−P of 86 Hz. These NMR properties are similar to other reported RhIII-monohydride complexes of similar structure.16 The measured pKa of the monohydride is 3.6 ± 0.3. Additionally, the 31P resonance for 1 shifts downfield as the pH is decreased, moving from 2.20 ppm at pH 8.0 to 8.1 ppm at pH 3.5. This shift is attributed to protonation of the carboxylates of the ligand backbone. Consistent with this assignment, the JRh−P coupling constant, 126 Hz, remains unchanged as the pH decreases, indicating that the ligand environment at the metal center is unchanged.16 The monohydride species was not observable within the protein scaffold due to the reduced stability of the scaffold at low 623

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pH. However, on the basis of the measured pKa of monohydride species of 1, the monohydride intermediate should be immediately deprotonated under catalytic conditions (pH 8.15), suggesting that the gain of catalytic function within the protein scaffold is not achieved by altering the pKa of the monohydride species. Consistent with that interpretation, only the RhI species is observable at pH 8.15 for both 1 and 2, Figure 2A,B. In comparing the reactivities of 1 and 2, they both react readily with H2, and their corresponding monohydride intermediates have low pKa values relative to catalytic conditions. The difference in reactivity is in the CO 2 hydrogenation step. Metalloenzyme 2 is capable of catalytically transferring a hydride to CO2, while 1 cannot. This suggests that interactions between the rhodium complex and the protein scaffold are facilitating the hydride transfer step. In support of this assertion, the catalytic activity of 2 decreases by 50% when the temperature is increased to 323 K, which is slightly below the TM estimated for 2. Additionally, catalytic activity is lost at 353 K where the protein is fully denatured, and precipitation occurs, Figure S1. The protein scaffold could be orienting CO2, thus facilitating interactions with the dihydride intermediate through hydrogen bonding with specific residues. However, on the basis of the 2:1 crystal structure, it is not obvious how the second coordination sphere contributes to the catalytic observations. Conserved arginine and histidine residues are proposed to serve this function in metal-dependent FDHs.7 The role of several arginine residues is under further investigation through mutagenesis and computational studies. The artificial metalloenzyme reported here constitutes a rare example of a synthetic inorganic complex gaining catalytic function upon incorporation into a protein scaffold without changes to the primary coordination sphere.29 One typical approach to create an artificial metalloenzyme is to incorporate an active molecular catalyst into a protein scaffold in order to control reactivity through changes to the environment around the metal30−32 or through changes to the ligand set of the metal.33,34 A fully active, bioidentical [FeFe]-hydrogenase can be matured from synthetically prepared, catalytically incompetent cofactors, but the protein scaffold enforces isomerization and a change in metal ligation, compared to the synthetically prepared cofactor, to achieve the catalytically active species.35 Uniquely, in the artificial metalloenzyme 2 reported here, the protein scaffold imparts catalytic activity onto an inactive, inorganic complex solely through secondary and outer coordination sphere effects. In summary, the covalent attachment of a rhodium bis(diphosphine) complex to a protein scaffold results in a gain of function generating an active metalloenzyme for CO2 hydrogenation. Structural characterization of this metalloenzyme and proposed catalytic intermediates suggests that the ligation and geometry of the metal center is not altered by attachment. However, the metalloenzyme can catalytically hydrogenate CO2, while the soluble complex in identical solution conditions cannot. The gain of function illustrates the profound influence that the extended environment around an active site can have on the reactivity even though the properties of the metal center in and out of the scaffold appear very similar. The gain of catalytic activity observed in this system exemplifies the importance of outer sphere interactions for catalytic function and their necessity in catalyst design. Current studies are focused on how the protein scaffold activates catalysis.

Research Article

ASSOCIATED CONTENT

S Supporting Information *

The Supporting Information is available free of charge on the ACS Publications website at DOI: 10.1021/acscatal.8b02615.

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Experimental methods, circular dichroism spectra, thermal denaturation curve, NMR data, kinetic plots, crystal structure data, and refinement statistics (PDF) X-ray structure validation report (PDF)

AUTHOR INFORMATION

Corresponding Author

*E-mail: [email protected]. ORCID

John C. Linehan: 0000-0001-8942-7163 Molly O’Hagan: 0000-0002-8344-8497 Present Address ⊥

Department of Chemistry and Biochemistry, Montana State University, Bozeman, MT 59717, U.S.A. Notes

The authors declare no competing financial interest.

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ACKNOWLEDGMENTS We thank Dr. Eric Walter for help with labeling studies. Research was funded by the Laboratory Directed Research and Development program at Pacific Northwest National Laboratory. The Pacific Northwest National Laboratory is operated by Battelle for the U.S. Department of Energy. This structural work is supported as part of the Biological and Electron Transfer and Catalysis (BETCy) EFRC, an Energy Frontier Research Center funded by the U.S. Department of Energy, Office of Science (DE-SC0012518) to J.W.P. and O.A.Z. This work was performed in part using the William R. Wiley Environmental Molecular Sciences Laboratory, a U.S. Department of Energy (DOE) national scientific user facility sponsored by the DOE’s Office of Biological and Environmental Research and located at the Pacific Northwest National Laboratory (PNNL). Use of the Stanford Synchrotron Radiation Lightsource, SLAC National Accelerator Laboratory, is supported by the U.S. Department of Energy, Office of Science, Office of Basic Energy Sciences under Contract No. DE-AC02-76SF00515. The SSRL Structural Molecular Biology Program is supported by the DOE Office of Biological and Environmental Research and by the National Institutes of Health, National Institute of General Medical Sciences (including P41GM103393).

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REFERENCES

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DOI: 10.1021/acscatal.8b02615 ACS Catal. 2019, 9, 620−625