From micelles to bicelles: Effect of the membrane on particulate methane monooxygenase activity

Abstract

Particulate methane monooxygenase (pMMO) is a copper-dependent integral membrane metalloenzyme that converts methane to methanol in methanotrophic bacteria. Studies of isolated pMMO have been hindered by loss of enzymatic activity upon its removal from the native membrane. To characterize pMMO in a membrane-like environment, we reconstituted pMMOs from Methylococcus (Mcc.) capsulatus (Bath) and Methylomicrobium (Mm.) alcaliphilum 20Z into bicelles. Reconstitution into bicelles recovers methane oxidation activity lost upon detergent solubilization and purification without substantial alterations to copper content or copper electronic structure, as observed by electron paramagnetic resonance (EPR) spectroscopy. These findings suggest that loss of pMMO activity upon isolation is due to removal from the membranes rather than caused by loss of the catalytic copper ions. A 2.7 Å resolution crystal structure of pMMO from Mm. alcaliphilum 20Z reveals a mononuclear copper center in the PmoB subunit and indicates that the transmembrane PmoC subunit may be conformationally flexible. Finally, results from extended X-ray absorption fine structure (EXAFS) analysis of pMMO from Mm. alcaliphilum 20Z were consistent with the observed monocopper center in the PmoB subunit. These results underscore the importance of studying membrane proteins in a membrane-like environment and provide valuable insight into pMMO function.

Introduction

Methanotrophic bacteria convert methane, the second most abundant greenhouse gas, to methanol in the first step of their metabolic pathway (1, 2). As the main methane sink in nature, these microorganisms are promising biological tools for methane remediation and biofuel production (3–6). Methanotrophs activate a 105 kcal/mol C-H bond in methane using metalloenzymes called methane monooxygenases (MMOs),³ which are classified as soluble or membrane-bound (particulate, pMMO) (7). pMMO is the predominant methane oxidation catalyst in nature but is less well-characterized (8). A detailed understanding of methane oxidation by pMMO has the potential to guide synthetic catalyst design and facilitate methanotroph engineering.

pMMO is a complex integral membrane enzyme that requires copper for activity (9–11). Crystal structures of pMMO from four different methanotrophs reveal a 300-kDa α₃β₃γ₃ trimer composed of the subunits PmoA, PmoB, and PmoC (11–14). PmoA and PmoC are integral membrane subunits, whereas PmoB consists of two periplasmic domains linked by two transmembrane helices. Present in all of these structures is a copper site at the N terminus of PmoB, with the N-terminal histidine of PmoB and two histidines from an HXH motif as ligands. This copper center, assigned as the active site (10), has been modeled with either one or two copper ions in the different structures. An additional PmoB monocopper site is found only in the structure of pMMO from Methylococcus (Mcc.) capsulatus (Bath) (12). The PmoC subunit houses a variable metal binding site that can be occupied by copper or zinc, depending on the crystallization conditions.

The presence of multiple subunits with variable metal content has complicated efforts to determine the nuclearity of the copper active site and the roles of the other observed metal centers. Moreover, all studies of pMMO have been hindered by significant decreases in enzymatic activity upon isolation from the membranes and solubilization with detergents (7). In some cases, activity appears to be completely abolished upon removal from the membranes. As a result, the physiological relevance of structural and spectroscopic studies of purified pMMO has been questioned, and alternative hypotheses for the active site have been proposed, largely based on the assumption that solubilization and purification of pMMO removes catalytically essential copper ions (15, 16).

An alternative possibility is that removal of pMMO from the membranes, rather than loss of copper, has deleterious effects on activity. Detergent micelles are frequently used for membrane protein characterization because of ease of use and compatibility with many experimental methods (17, 18). However, detergent micelles lack the structure and pressure provided by lipid bilayers and can cause instability and loss of function (19, 20). Membrane mimetics provide a way to study membrane proteins in more native-like environments. In many cases, addition of lipids or use of these mimetics has restored functional activity to purified membrane proteins (21–24). In particular, bicelles, discoidal lipid bilayers surrounded by detergent, have been used to characterize and crystallize a range of membrane proteins (25–28).

To address the hypothesis that pMMO inactivation upon solubilization is due to removal from the native membranes, we have reconstituted purified pMMO from Methylomicrobium (Mm.) alcaliphilum 20Z (20Z-pMMO) and Mcc. capsulatus (Bath) (Bath-pMMO) into bicelles. Bicelle reconstitution recovers the methane oxidation activity of both pMMOs without addition of exogenous copper ions or substantial alteration in the copper sites, as observed by electron paramagnetic spectroscopy (EPR). A crystal structure of 20Z-pMMO provides some insight into how solubilization might affect protein stability. Finally, extended X-ray absorption fine structure (EXAFS) analysis of 20Z-pMMO does not indicate the presence of the short copper–copper interaction observed in previous samples, prompting further investigation of the active site nuclearity.

Results

Recovery of [¹³C]methane oxidation activity by bicelle reconstitution

To systematically investigate loss of pMMO activity, methane oxidation activity was measured for as-isolated, solubilized, purified, and bicelle-reconstituted pMMO samples. pMMO activity assays are typically performed using either NADH or duroquinol as a reductant. Duroquinol can directly reduce pMMO, whereas a type 2 NADH dehydrogenase (NDH-2) likely oxidizes NADH and reduces quinones for subsequent electron transfer to pMMO (9). Solubilization with the detergent dodecyl maltoside (DDM) separates pMMO from the membranes (solubilized pMMO), which abrogates NADH-driven activity (Fig. 1). Solubilized pMMO was then reconstituted in bicelles to mimic the lipid bilayer and to investigate membrane-dependent activity loss. Methane oxidation activity was measured for as-isolated membranes, solubilized and purified pMMO in detergent (DDM), and bicelle (3% (w/v) DMPC-CHAPSO) reconstituted pMMO using both reductants (Fig. 1). Because of [¹²C]methanol contamination in many buffers and reagents, a new activity assay was developed in which conversion of [¹³C]methane to [¹³C]methanol is detected via GC-MS.

Figure 1.

Methane oxidation activity of pMMOs in native membranes, detergent, and bicelles. A and B, activity values are shown for Bath-pMMO (A) and 20Z-pMMO (B) in as-isolated membranes, solubilized in detergent, solubilized in bicelles, purified in detergent, and purified in bicelles using both duroquinol (gray) and NADH (white) as reductants. Error bars represent standard deviations of three measurements, and the black and white dots represent the individual measurements.

Methane oxidation activities for Bath-pMMO and 20Z-pMMO were measured at 30 °C after 5 min because of solidification of bicelles at higher temperatures and longer incubation times. NADH-driven activity (41.1 ± 1.7 and 14.5 ± 1.2 nmol [¹³C]methanol mg⁻¹ protein for Bath-pMMO and 20Z-pMMO, respectively) is abolished upon solubilization and purification (Fig. 1 and Table S1). For membrane-bound and solubilized samples, the activity measured using duroquinol was significantly lower than the NADH-driven activity for Bath-pMMO and not detected for 20Z-pMMO (Fig. 1). For both pMMOs, reconstitution into bicelles recovers the methane oxidation activity of solubilized and purified samples using duroquinol as a reductant (Fig. 1 and Table S1). However, NADH-driven activity is only restored for Bath-pMMO. It may be that an NDH-2 or other components of the electron transport chain responsible for NADH-dependent methane oxidation are not properly reassembled after solubilization and reconstitution of 20Z-pMMO.

Notably, for both pMMOs, duroquinol-driven activity is significantly higher for bicelle-reconstituted samples than for as-isolated membranes and is comparable with NADH-driven activity in membranes (Fig. 1 and Table S1). The different properties of phosphatidylcholine (PC), the main lipid in DMPC bicelles, and phosphatidylethanolamine (PE), the predominant phospholipid found in these methanotrophs (29–31), provide a possible explanation for this observation. The amine head group of PC is less polar than that of PE and may increase the solubility and access of duroquinol as well as O₂ and methane. Additionally, DMPC is composed of saturated 14:0 PC, whereas methanotroph PEs are primarily composed of a saturated and unsaturated mixture of 16:0 and 16:1 PE. The various head groups and acyl chain compositions can affect lipid packing, membrane fluidity, and even the structure of membrane proteins (32). Finally, in as-isolated membranes, it is possible that the native quinones occupy the binding site duroquinol needs to access to reduce pMMO. Taken together, these results indicate that solubilized pMMOs are not irreversibly inactivated. Interestingly, solubilized or purified pMMO samples were reconstituted in bicelles without the addition of copper, suggesting that bicelles alone are responsible for the recovered activity.

Effect of bicelle reconstitution on pMMO copper centers

To further investigate the relationship between bicelle reconstitution and the pMMO copper sites, the copper concentrations of pMMO samples in as-isolated membranes, detergent, and bicelles were measured using inductively coupled plasma optical emission spectroscopy. The presence of approximately three copper ions per protomer in purified Bath-pMMO (Fig. 2 and Table S2) is consistent with previous studies (12). Purified 20Z-pMMO contains ∼2.7 eq of copper per protomer (Table S2). The copper contents of the native membranes and solubilized pMMOs are batch-dependent, accounting for the variability in copper stoichiometry values for these samples. Loss of some adventitiously bound copper is also typically observed during solubilization and purification (11).

Figure 2.

Copper content of pMMOs in native membranes, detergent, and bicelles. A and B, copper stoichiometry of Bath-pMMO (A) and 20Z-pMMO (B) in as-isolated membranes, solubilized in detergent, solubilized in bicelles, purified in detergent, and purified in bicelles. Error bars represent standard deviations of three measurements, and the black dots represent individual measurements.

The copper stoichiometry does not change between pMMO samples in detergent and in bicelles (Fig. 2 and Table S2). This observation, in conjunction with the recovered activity, indicates that the catalytically essential copper ions are still present in detergent-solubilized pMMO samples. The differences in activity between as-isolated membranes, detergent-solubilized pMMO, and bicelle-reconstituted pMMO therefore cannot be attributed to changes in copper content. Consequently, the membrane, and not copper depletion, is a crucial factor contributing to activity loss upon solubilization.

To directly assess the Cu²⁺ electronic and geometric structure through the bicelle reconstitution process, we collected EPR spectra of Bath-pMMO and 20Z-pMMO before and after bicelle reconstitution (Fig. 3). A previous EPR analysis of purified Bath-pMMO revealed two distinct type 2 Cu²⁺ signatures (33). The bicelle-reconstituted Bath-pMMO exhibits the same Cu²⁺ EPR spectrum as the purified Bath-pMMO and is simulated with the same parameters as reported previously. However, the bicelle-reconstituted enzyme contains slightly more Cu²⁺ per protomer than the purified sample. Consequently, some of the Cu²⁺ observed in the bicelle-reconstituted sample is Cu¹⁺ in the purified sample and oxidizes to Cu²⁺ during the reconstitution procedure.

Figure 3.

X-band EPR spectra of Bath-pMMO and 20Z-pMMO. Spectra of pMMO purified in detergent (gray, dotted lines) and after bicelle reconstitution (black, solid lines) are normalized to protein concentration. For purified and bicelle-reconstituted Bath-pMMO, EPR intensity was increased 3-fold for ease of comparison. Red solid lines depict simulations using the (very slightly adjusted) previously defined Cu²⁺ spin Hamiltonian parameters (33): equal contributions of (His)₂-Cu²⁺ (g_‖ = 2.235, g_⊥ = 2.047, A_‖ = 585 MHz, A_⊥ = 65 MHz) and (His, Amine)-Cu²⁺ (g_‖ = 2.295, g_⊥ = 2.047, A_‖ = 450 MHz, A_⊥ = 40 MHz). Collection conditions were as follows: 9.36–9.37 GHz microwave frequency, 20 K temperature, 160 ms time constant, 16 G modulation amplitude, 60-s scans, average of at least 10 scans.

The purified 20Z-pMMO EPR spectrum exhibits the Cu²⁺ spectrum seen in both forms of Bath-pMMO as well as a small contribution from additional Cu²⁺ resonance (Fig. 3B and Table S2), suggesting adventitious Cu²⁺ binding to 20Z-pMMO in a site either unoccupied or containing Cu¹⁺ in Bath-pMMO. Similar to Bath-pMMO, incorporation of 20Z-pMMO into bicelles oxidizes some Cu¹⁺ to Cu²⁺, as evidenced by the slightly altered g_⊥ region and increased amount of Cu²⁺ per protomer (Table S2), but the signal is otherwise the same as observed for the purified sample.

Importantly, the EPR spectra of both pMMOs show that the Cu²⁺ ligation is not substantially altered by the bicelle incorporation procedure. Therefore, the appreciable recovery of pMMO activity upon insertion of Bath-pMMO into the bicelle is not due to differences in the active site copper structure, consistent with the notion that the membrane environment plays a critical role in modulating activity.

Crystal structure of 20Z-pMMO

To further characterize 20Z-pMMO, a crystal structure was determined to 2.7 Å resolution (Table 1). The protein was purified in the presence of DDM, exchanged into the detergent Cymal-5, and then crystallized with ammonium sulfate as the precipitant. Varying the concentration of this precipitant was crucial for obtaining well-diffracting crystals. The 20Z-pMMO structure exhibits a similar overall architecture to Bath-pMMO, with an α₃β₃γ₃ trimeric structure. Unlike previous pMMO structures (11–14), there is a single protomer in the asymmetric unit (Fig. 4A).

Table 1.

Data collection and refinement statistics for 20Z-pMMO

Values in parentheses refer to the highest-resolution shell.

	20Z-pMMO	20Z-pMMO (copper anomalous)
Data collection
Space group	P63	P63
Cell dimensions
a, b, c	143.84, 143.84, 146.15
Resolution (Å)	2.70	3.00
Wavelength (Å)	1.03329	1.37760
Rpim	0.025 (0.220)	0.022 (1.32)
Rmeas	0.07 (0.588)	0.06 (0.337)
CC½	0.999 (0.953)	0.998 (0.984)
I/σI	40.6 (2.5)	113.5 (11.2)
Completeness (%)	99.1 (92)	99.7 (97.7)
Redundancy	9.0 (6.4)	6.9 (5.7)
Anisotropy correction
Truncation limit (Å)	3.1, 3.1, 2.7
Completeness (%)	74.9
Refinement
No. of reflections	35,187
Rwork/Rfree	0.2133/0.267
Average B-factor (Å2)	36.49
Root mean square deviations
Bond lengths (Å)	0.010
Bond angles (°)	1.313
Ramachandran favored (%)	87.39
Ramachandran allowed (%)	10.64

Figure 4.

Crystal structure of 20Z-pMMO. A, the 20Z-pMMO protomer with PmoB, PmoA, and PmoC shown in gray, salmon, and purple, respectively. The copper ion is depicted as a teal sphere. B, superposition of the 20Z-pMMO PmoC subunit (purple) with the Bath-pMMO PmoC subunit (PDB code 3RGB, yellow). The zinc ion in the Bath-pMMO variable metal-binding site is shown as a sphere (gray) and is coordinated by two histidines and an aspartic acid. This region is disordered in 20Z-pMMO. C, the site of the monocopper center in Bath-pMMO is unoccupied in 20Z-pMMO. D, the 20Z-pMMO copper site. The copper anomalous difference density map (magenta, 20 σ) is superimposed on the 2F_o-F_c map (gray, 1.0 σ).

Despite the overall structural similarity, the PmoC subunit of 20Z-pMMO is significantly disordered compared with the previous structures (11–14) (Fig. 4B). Electron density is not observed for 60% of the PmoC subunit, including residues 1–89, 123–156, and 193–218. These disordered regions include the variable metal binding site (Asp-128, His-132, and His-145) and surrounding residues. This significant disorder may result from destabilization of PmoC in detergent and could be related to the complete loss of activity upon detergent solubilization and purification (Fig. 1B). PmoC, at least in 20Z-pMMO, is thus more flexible than suggested by previous structures.

The metal binding sites of 20Z-pMMO also differ from those observed in previous pMMO structures (11–14). In the Bath-pMMO PmoB subunit, there is a monocopper site coordinated by His-48 and His-72 (12). Although both residues are conserved in 20Z-pMMO, electron density attributable to copper or any other metal ion is not present (Fig. 4C). It is unclear why this site is only occupied in Bath-pMMO, but the metal binding residues are not conserved in all pMMOs, with His-48 substituted by Asn and Gln in type II methanotrophs, indicating that this metal center is not essential for methane oxidation. The PmoB subunit also contains a bound copper that is coordinated by residues His-33, His-137, and His-139 and has been assigned as the active site. In some pMMO structures, this site has been modeled with two copper ions, including Bath-pMMO (12–14). The dicopper site model is based on EXAFS data that consistently indicate the presence of a short copper–copper distance as well as the measured copper stoichiometry upon purification (10, 11, 13, 14, 34). However, in other structures, the site has been modeled with a single copper ion (11, 14).

In the 20Z-pMMO structure, this PmoB site is also best modeled with one copper ion (Fig. 4D). The site is square planar with copper–nitrogen distances of 2.1 Å for the His-137 δN, 2.1 Å for the His-139 ϵN, 2.5 Å for the His-33 δN, and 1.9 Å for the N-terminal nitrogen of His-33. The electron density for His-33 is not as well-defined as that for other two histidine residues. Interestingly, very strong additional electron density is observed for PmoB residue Lys-155 in PmoB appended to the side-chain ζN atom. We could not conclusively model this density, but it could potentially arise from posttranslational modification of this residue.

XANES and EXAFS analysis of 20Z-pMMO

The copper X-ray absorption near edge structure (XANES) spectra measured for 20Z-pMMO indicate a mixed Cu(I) and Cu(II) metal environment. A subtle transition, observed at 8978.8 eV (Fig. 5A) is consistent with the forbidden 1s → 3d transition for Cu(II) (35). Additional edge transitions, observed at 8983 and 8986.3 eV and illustrated in the first derivative of the edge at 8982.3 and 8985.5 eV in Fig. 5A, inset, are characteristic of the 1s → 4p transitions often observed for systems containing a mixture of Cu(I) and Cu(II) (35).

Figure 5.

XANES and EXAFS analysis of 20Z-pMMO. A, copper XANES spectra for 20Z-pMMO. Inset, the first derivative of near edge and edge features is displayed to more clearly highlight the features. B, raw copper EXAFS for 20Z-pMMO. Simulations were fit using a standard conservative approach that follows rules governing both spectral resolution relative to acceptable intraligand scattering interaction bond lengths and acceptable bond lengths (55). C, Fourier transform of the EXAFS. Raw unfiltered data are shown in black, and the best fit simulations are shown in gray. EXAFS were fit over a k range of 1.0–12.85 Å⁻¹.

Analysis of the copper EXAFS spectra for 20Z-pMMO suggest a mononuclear copper ligand environment constructed by only oxygen and nitrogen within the first ligand sphere (Fig. 5B). Simulations of copper–oxygen/nitrogen nearest neighbor ligand scattering suggest a disordered ligand environment composed of approximately 2.5 to 3.5 oxygen/nitrogen ligands at an average bond length of 1.96 Å (Table 2). Inclusion of a direct copper–copper scattering vector was not justified in our simulations. Long-range scattering could be simulated using low Z (carbon/nitrogen) scattering at bond lengths of 2.97, 3.36, and 3.97 Å, reminiscent of patterns observed because of imidazole scattering interactions from coordinated histidines (36). In support of imidazole scattering, the pronounced camelback feature at 4 Å⁻¹, characteristic of metal–histidine ligation (37), is also observed.

Table 2.

Summary of the best fit copper EXAFS simulations for the 20Z-pMMO sample

EXAFS were fit over the k range of 1.0–12.85 Å⁻¹ for a spectral resolution of 0.13 Å.

Nearest neighbor ligand environment				Long-range ligand environment				F′e
Atoma	R (Å)b	CNc	σ2d	Atoma	R (Å)b	CNc	σ2d
O/N	1.96	2.5	5.23	C	2.97	1.5	3.73	0.4
				C	3.36	0.75	4.25
				C	3.96	3.0	4.22

^a Scattering atoms: O, oxygen, N, nitrogen; C, carbon.

^b Average metal–ligand bond length from two independent samples.

^c Average metal–ligand coordination number from two independent samples.

^d Average Debye–Waller factor in Ų × 10³ from two independent samples.

^e Number of degrees of freedom weighted mean square deviation between data and fit.

Discussion

The recovery of methane oxidation activity upon pMMO reconstitution into bicelles underscores the importance of studying membrane proteins in native-like environments. Although studying membrane proteins in a membrane context seems obvious, detergent micelles are still typically used instead. Besides their amphipathic nature, detergent micelles lack important lipid bilayer characteristics that provide structural support (19). Reconstitution of pMMO into bicelles restores the methane oxidation activity of inactive detergent-solubilized pMMOs close to levels measured for membrane-bound pMMO (Fig. 1). The copper stoichiometries and EPR spectroscopic features are nearly identical for inactive detergent-solubilized and active bicelle-reconstituted pMMO samples and are consistent with previous observations (12, 33).

These data indicate that the copper centers detected in detergent-solubilized pMMO are functionally relevant. In previous pMMO crystal structures, one to three copper ions were modeled per protomer, found only in the PmoB and PmoC metal centers, and only the PmoB site coordinated by His-33, His-137, and His-139 consistently houses copper ions (11–14). Chan and Yu (15) and Chan and co-workers (16) have proposed that Bath-pMMO instead contains ∼15 copper ions, including a tricopper active site in PmoA and six to seven Cu¹⁺ ions bound to the C terminus of PmoB, and have suggested that copper loss from these sites upon membrane solubilization is responsible for the reduced activity of purified Bath-pMMO. However, the recovered activity of bicelle-reconstituted pMMO samples indicates that large numbers of essential copper ions are not lost during isolation from the membranes.

The crystal structure of 20Z-pMMO provides some insight into how removal from the membrane could affect activity. PmoC is largely disordered, suggesting destabilization upon solubilization and resultant activity loss. PmoB only contains two transmembrane helices, and PmoA is sandwiched between PmoB and PmoC, features that may contribute to their structural stability in detergent micelles. In contrast, only the PmoC helices near PmoA are ordered (Fig. 4A), whereas the disordered regions are exposed to the lipid membrane and perhaps more susceptible to perturbations upon reconstitution into detergent micelles. Without lateral pressure or specific lipid binding, PmoC may be structurally less stable in micelles. PmoC is positioned directly adjacent to the proposed PmoB active site and could be involved in stabilization of the active site or copper binding that may be essential for activity. In addition, for a hydrocarbon monooxygenase homolog of pMMO, mutation of the PmoC metal binding residues reduces activity, suggesting an important functional role (38). Previous efforts have mainly focused on characterizing perturbations in PmoB to explain activity loss. Some of this attention should be shifted to understanding how the transmembrane subunits, particularly PmoC, play an essential role in methane oxidation.

Finally, a mononuclear copper active site remains a viable possibility (6, 8). The 20Z-pMMO PmoB site is best modeled as monocopper (Fig. 4). Additionally, the short copper–copper distance detected in the EXAFS analysis of other pMMOs (11, 14, 35) is not present in 20Z-pMMO (Fig. 5). Its absence in 20Z-pMMO could be due to lower protein concentrations, a heterogeneous distribution of copper–copper vectors in the samples that cancel out the overall signal, or even the reduced presence of other copper contaminant proteins that could contribute to the observed feature. This result is consistent with a recent quantum refinement of the Bath-pMMO PmoB copper site (39). Most relevant to a pMMO monocopper active site are the lytic polysaccharide monooxygenases (LPMOs), which utilize a monocopper active site for oxidative cleavage of glyosidic bonds. Both the PmoB copper site and the LPMO active site contain a histidine brace metal-binding motif. However, LPMOs lack a third histidine ligand and additional metal binding sites. In addition, some contain a methylated histidine ligand. Moreover, the substrates of pMMO and LPMO are drastically different (40–42). Overall, studying pMMO in a membrane-bound context has validated past characterizations and provides new insights into the importance of the PmoC subunit and the nature of the active site. It will be important to continue this approach in future studies of pMMO activity and mechanism.

Experimental procedures

Methanotroph cell growth

Mm. alcaliphilum 20Z was cultured as described previously (43, 44). Briefly, cells were grown in 1× modified nitrate mineral salts medium, 0.5 m NaCl, 2.3 mm phosphate buffer, and 50 mm carbonate buffer (pH 9.5) supplemented with 40 μm CuSO₄·H₂O and trace elements solution under a 1:3 methane-to-air ratio in 12 liters bioreactors. Mcc. capsulatus (Bath) cells were grown in 1× nitrate mineral salts medium and 3.9 mm phosphate buffer (pH 6.8) supplemented with 50 μm CuSO₄·H₂O, 40 μm iron NaFe(III)EDTA, and trace element solution under a 1:4 methane-to-air ration in 12 liters of bioreactors (12). All bioreactor cell growths were harvested at an OD₆₀₀ of 8–10 and centrifuged at 8,000 × g for 30 min. Cell pellets were flash-frozen in liquid nitrogen and stored at −80 °C for future use.

Membrane isolation

Mm. alcaliphilum 20Z cell pellets (10 g) were resuspended in 100 ml of 25 mm PIPES and 500 mm NaCl (pH 7) supplemented with EDTA-free protease inhibitor tablets (Roche). The cells were manually stirred for resuspension on ice. The cell resuspension was sonicated at 4 °C for 1.5 min with an on/off interval of 1 s/3 s at 25% amplitude and centrifuged at 8,000 × g for 1 h to remove cell debris. The supernatant was centrifuged at 100,000 × g for 1 h to isolate the pelleted membranes containing pMMO. The membrane pellet was washed twice with a Dounce homogenizer in 25 mm PIPES and 250 mm NaCl (pH 7). 1-ml aliquots of pMMO-containing membranes at total protein concentrations of 10 mg/ml (measured by Bio-Rad DC assay using BSA as a standard) were flash-frozen in liquid nitrogen and stored at −80 °C. Mcc. capsulatus (Bath) membranes were isolated as described previously (45).

pMMO purification and bicelle reconstitution

Membranes were solubilized using 1.2 mg of DDM (Anatrace) per 1 mg of crude protein at 4 °C for 1 h. The solubilized protein was centrifuged at 100,000 × g for 1 h, and the supernatant was collected for purification. Solubilized 20Z-pMMO was buffer-exchanged into 25 mm PIPES, 50 mm NaCl (pH 7), and 0.02% (w/v) DDM using a 100-kDa molecular mass cutoff Amicon (Millipore). 20Z-pMMO was purified using a 15Q anion exchange column (GE Healthcare) and eluted using a 50–800 mm NaCl gradient (Figs. S1 and S2). Solubilized Bath-pMMO was concentrated to 1 ml using a 100-kDa molecular mass cutoff Amicon and loaded onto a 120-ml Superdex 200 size exclusion column (Fig. S3). All eluted pMMOs were concentrated using a 100-kDa molecular mass cutoff Amicon to 10 mg/ml in 25 mm PIPES, 250 mm NaCl (pH 7), and 0.02% (w/v) DDM. Freshly solubilized or purified pMMO at 10 mg/ml was reconstituted with a 30% (w/v) DMPC:CHAPSO 2.8:1 bicelle solution (Molecular Dimensions) using a 4:1 volumetric ratio and incubated on ice for at least 30 min to prepare pMMO samples at 8 mg/ml reconstituted in 6% (w/v) bicelles. The copper concentration was measured by inductively coupled plasma optical emission spectroscopy at the Quantitative Bio-element Imaging Center at Northwestern University.

[¹³C]Methane oxidation activity assay

To measure the methane oxidation activity of membrane-bound, solubilized, purified, and 6% (w/v) bicelle-reconstituted pMMOs, samples were diluted to 4 mg/ml (or 3% (w/v) bicelles) in 100-μl reactions consisting of reductant (280 μm NADH (Sigma-Aldrich) or excess duroquinol) in 2-ml screw-top vials with septa tops (Agilent). A 1-ml volume of headspace gas was withdrawn and replaced with 1.5 ml of [¹³C]methane (Sigma-Aldrich). All reactions were performed at 30 °C (bicelle samples solidify at 45 °C, the temperature typically used for Bath-pMMO activity assays). pMMO reconstituted in 3% or 1.5% (w/v) bicelles gave the highest activity, which decreased with lower bicelle concentrations (Fig. S4). Reactions were incubated at 30 °C and 200 rpm for 5 min, put on ice for 5 min, and then quenched with 500 μl of chloroform spiked with 1 mm dichloromethane. The reaction was vortexed at 2,000 rpm for 10 min and centrifuged at 2,000 × g for 30 min to separate precipitate from the chloroform mixture. 2.5 μl of sample was injected onto a PoraBOND Q column (25 m × 250 μm × 3 μm) on an Agilent 7890B/5977A MSD GC/MS instrument with a split ratio of 10:1. The GC was maintained under a constant flow of 1.2 ml/min of helium gas. The initial oven temperature was maintained at 80 °C for 3.5 min, followed by an increase of 50 °C/min to 150 °C and held for 1.5 min. A second ramp rate of 15 °C/min was used to reach the final temperature of 300 °C, held for 1 min. The mass spectrometer was maintained under an ion source temperature of 230 °C, quad temperature of 150 °C, 70 eV, and a detector voltage of 2,999 V. Masses 31, 33, and 49 were monitored for detection of [¹²C]methanol, [¹³C]methanol and dichloromethane (dwell times of 10 ms, 100 ms, and 10 ms, respectively). The [¹³C]methanol peak area was integrated, quantified from a standard calibration curve, and normalized to the concentration of the internal standard dichloromethane. The lower limit of detection was determined to be 10 μm [¹³C]methanol, and a stringent cutoff for minimum concentration was set at 30 μm. Methane oxidation activity values using [¹³C]methanol detection by GC-MS compared with [¹²C]methanol detection using the GC-flame ionization detector (FID) are shown in Table S3.

EPR spectroscopy

EPR samples were prepared by aliquoting 100 μm (DDM samples) or 80 μm pMMO (bicelle samples) in 25 mm PIPES, 250 mm NaCl (pH 7), and 0.02% (w/v) DDM or 6% (w/v) bicelles into Wilmad quartz EPR tubes (Sigma-Aldrich). Measurements were collected on a continuous wave X-band Bruker ESP-300 spectrometer using a liquid helium flow Oxford Instruments ESR-900 cryostat. Spectra were corrected for background resonance by subtraction of a spectrum of 50 mm Tris (pH 8.0), 150 mm NaCl collected under the same conditions. Cu²⁺ spin quantitation was performed by double integral area comparison of pMMO spectra to Cu²⁺–EDTA standards containing 25–500 μm Cu²⁺. All EPR simulations were performed using EasySpin (46).

Crystallization and structural determination of 20Z-pMMO

Purified pMMO in 0.02% (w/v) DDM was buffer-exchanged into 0.12% (w/v) Cymal-5 using a 100-kDa molecular mass cutoff Amicon. pMMO crystals were obtained from sitting drops containing 1 μl of 10 mg/ml protein in 25 mm PIPES, 250 mm NaCl (pH 7), 0.12% (w/v) Cymal-5, 1 μl of 2.8 m AmSO₄, and 0.2 m MES (pH 6). Crystals were harvested in saturated LiSO₄ cryoprotectant solution and flash-frozen in liquid nitrogen.

Crystals were screened for diffraction at the LS-CAT and GM/CA-CAT beamlines at the Advanced Photon Source at Argonne National Laboratory. Datasets were processed using HKL2000 (47) (Table 1). Anisotropic processing using the UCLA anisotropy server (48) was found to improve the electron density maps. Phenix (49) was used for molecular replacement with the Bath-pMMO coordinates as a starting model (PDB code 3RGB) to solve the structure of 20Z-pMMO. 20Z-pMMO has 72%:78%:77% identity to the Bath-pMMO PmoB, PmoA, and PmoC subunits, respectively. Structure modeling and refinement were performed using Coot (50) and Phenix, and model quality was assessed using MolProbity (51). The final model for the 20Z-pMMO structure includes PmoB residues 33–414; PmoC residues 90–122, 157–192, and 219–250; PmoA residues 4–244; one copper ion; and two Cymal-5 molecules.

X-ray absorption spectroscopy

Purified 20Z-pMMO samples were concentrated to 385 μm using a 100-kDa molecular mass cutoff Amicon centrifugal concentrator and resuspended in 30% (v/v) glycerol. The copper concentration of the 20Z pMMO samples was 732 μm. These samples were loaded into Lucite XAS cells wrapped with Kapton tape, flash-frozen in liquid nitrogen, and stored at −80 °C. XAS data were collected at the Stanford Synchrotron Radiation Lightsource on beamline 9-3, equipped with a Si[220] double-crystal monochromator that contains an upstream mirror used for focusing and harmonic rejection. Fluorescence spectra were collected using a 100-element Ge solid-state Canberra detector. During data collection, the Oxford Instruments continuous-flow liquid helium cryostat was stabilized at 10 K. Copper excitation data were collected using a 6-μm nickel Lytle filter and solar slits placed between cryostat and detector to reduce scattering fluorescence. XAS spectra were measured using 5 eV steps in the pre-edge region (8,750–8,960 eV), 0.25 eV steps in the edge region (8,986–9,050 eV), and 0.05 Å⁻¹ increments in the EXAFS region (to k = 13.3 Å⁻¹), integrating from 1 to 25 s in a k³ weighted manner for a total scan length of ∼40 min. A copper foil spectrum was collected simultaneously with each protein spectrum for real-time spectral energy calibration, with an assigned first inflection point for the copper foil spectrum at 8,980.3 eV. Spectra were closely monitored for any photodamage, and slight photoreduction was observed. To diminish the extent and impact of photoreduction, six individual spectra were collected at unique positions on the sample surface, following a matrix positioning grid to ensure a new radiation exposure surface, and only the initial exposure spectrum at each position was used during overall data analysis. Spectra were collected on duplicate independent samples, and data presented in this report represent the average of six scans.

XAS spectra were processed and analyzed using the EXAFSPAK program suite written for Macintosh OS-X (52)⁴ integrated with the Feff v8 software (53) for theoretical model generation. EXAFS fitting analysis was performed on raw/unfiltered data. Single scattering models were calculated for oxygen, nitrogen, sulfur, copper, and carbon coordination to simulate possible copper ligand environments, with values for the scale factors (Sc) and E₀ calibrated by previous fittings of characterized Cu(I)/Cu(II) crystallographic copper model compounds (35). Standard criteria for judging the best-fit EXAFS simulations included a reasonable Debye–Waller factor for the fit (σ² < 0.006 Ų) (54); the spectral resolution of the data, calculated based on the energy range extent of usable data (55); and the lowest mean square deviation between data and fit width, corrected for the number of degrees of freedom (F′) (55). During the standard criteria simulations, only the bond length and Debye–Waller factor were allowed to vary for each ligand environment. A dimensionless Sc = 1 and E₀ values of −12, −14, and −16 eV were used for Cu(I,II)-C/N/O, -S, and -Cu theoretical model calibrations, respectively, during simulations (35).

Author contributions

S. Y. R., M. O. R., and A. C. R. conceptualization; S. Y. R., M. O. R., Y. W. D., S. B., T. J. L., and J. D. H. data curation; S. Y. R., M. O. R., Y. W. D., S. B., T. L. S., and B. M. H. formal analysis; S. Y. R., M. O. R., Y. W. D., S. B., T. L. S., B. M. H., and A. C. R. validation; S. Y. R., M. O. R., Y. W. D., S. B., T. J. L., J. D. H., T. L. S., and A. C. R. investigation; S. Y. R., M. O. R., S. B., and A. C. R. visualization; S. Y. R., M. O. R., Y. W. D., S. B., T. J. L., J. D. H., T. L. S., and A. C. R. methodology; S. Y. R., M. O. R., and A. C. R. writing-original draft; S. Y. R., M. O. R., S. B., T. L. S., B. M. H., and A. C. R. writing-review and editing; T. L. S., B. M. H., and A. C. R. supervision; T. L. S., B. M. H., and A. C. R. project administration; A. C. R. resources; A. C. R. funding acquisition.