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J Phys Chem B
J Phys Chem B
jp
jpcbfk
The Journal of Physical Chemistry. B
1520-6106
1520-5207
American Chemical Society

39231533
10.1021/acs.jpcb.4c03676
Article
Tracking Disordered Extracellular Domains of Membrane Proteins in the Cell with Cu(II)-Based Spin Labels
Meron Shelly †
Peleg Shahaf †
Shenberger Yulia
Hofmann Lukas
Gevorkyan-Airapetov Lada
https://orcid.org/0000-0002-1741-6892
Ruthstein Sharon *
The Department of Chemistry and the Institute of Nanotechnology and Advanced Materials, Faculty of Exact Sciences, Bar-Ilan University, Ramat-Gan 529002, Israel
* Email: Sharon.ruthstein@biu.ac.il.
04 09 2024
19 09 2024
128 37 89088914
03 06 2024
19 08 2024
30 07 2024
© 2024 The Authors. Published by American Chemical Society
2024
The Authors
https://creativecommons.org/licenses/by-nc-nd/4.0/ Permits non-commercial access and re-use, provided that author attribution and integrity are maintained; but does not permit creation of adaptations or other derivative works (https://creativecommons.org/licenses/by-nc-nd/4.0/).

In-cell electron paramagnetic resonance (EPR) spectroscopy experiments provide high-resolution data about conformational changes of proteins within the cell. However, one of the limitations of EPR is the requisite of stable paramagnetic centers in a reducing environment. We recently showed that histidine-rich sites in proteins hold a high affinity to Cu(II) ions complexed with a chelator. Using a chelator prevents the reduction of Cu(II) ions. Moreover, this spin-labeling methodology can be performed within the native cellular environment on any overexpressed protein without protein purification and delivery to the cell. Herein, we use this novel methodology to gain spatial information on the extracellular domain of the human copper transporter, hCtr1. Limited structural information on the transmembrane domain of the human Ctr1 (hCtr1) was obtained using X-ray crystallography and cryo-EM. However, these structures are missing information on the disordered extracellular domains of hCtr1. Extracellular domains are sensing or interacting with the environment outside of the cell and therefore play an essential role in any transmembrane protein. Especially in hCtr1, the extracellular domain functions as a gating mechanism for copper ions. Here, we performed EPR experiments revealing structural information about the extracellular N-terminal domain of the full-length hCtr1 in vitro and in situ in insect cells and cell membrane fragments. The comparison revealed that the extracellular domains of the in situ and native membrane hCtr1 are further apart than the structure of the purified protein. These method-related differences highlight the significance of studying membrane proteins in their native environment.

Israel Science Foundation 10.13039/501100003977 212/22 document-id-old-9jp4c03676
document-id-new-14jp4c03676
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pmcIntroduction

Electron paramagnetic resonance (EPR) spectroscopy has emerged as an excellent tool for gaining precise structural information on complex biological systems.1−4 However, targeting proteins within their native environment using EPR spectroscopy remains a challenging task.5 The future of structural biology lies in approaches that address proteins in their native environment. Hence, advances depend on our ability to integrate the entire cellular environment with all of its complexity. The advantages of EPR, over other methods, include higher sensitivity that allows for precise tracking of minor conformational changes in a targeted biomolecule, being unlimited to the size and complexity of the biological systems and its environment. In addition, EPR can target biomolecules present at concentrations as low as the micromolar range.2,6 Yet, EPR spectroscopy requires paramagnetic centers, a need that presents several challenges to use this technique for in-cell experiments. The first challenge is that spin labels must be stable in a reducing environment. To overcome this challenge, site-directed EPR spin labels commonly used for in-cell applications are Gd(III)-7 and trityl-based spin labels,8−11 which address this concern. A second limitation linked to the need for paramagnetic centers is that often the spin-labeling procedure is performed outside the cells. These spin-labeled proteins are then delivered into the cell. Protein transformation limits the size of the biomolecules of interest as well as the cellular system used, limited by how much the cell membrane can be distorted. The Saxena group has shown that Cu(II)-based distance measurements using the double-histidine (dHis) motif by pulsed EPR present an attractive strategy to obtain exact, narrow distance distributions that can be easily related to the protein backbone structure.12,13 In this approach, two strategically placed His residues are used to create a Cu(II) binding site. The Cu(II) ion is introduced as a complex with a chelating agent, which improves binding selectivity to the dHis site.14,15 We recently introduced this approach for in-cell spin labeling, where the Cu(II)–nitrilamino acid (NTA) complex was used to target conformational changes of proteins that are overexpressed with the dHis site in the cellular conditions.16 Spin labeling is performed directly in the cellular environment, thus omitting the need to purify the protein and deliver the labeled entity into the cells. Herein, we use this approach to gain structural information about the extracellular domain of the human copper transporter, hCtr1.

Following conformational changes in the extracellular and intracellular domains of ligand and ion transporters is essential to resolve their gating mechanism. These domains are usually disordered, which complicates the analysis of the conformational changes that these domains undergo upon ligand-ion binding and transfer mechanism. The Cafiso and Pliotas groups have shown that EPR spectroscopy can follow in situ conformational changes in the extracellular domain of membrane proteins in their native environment.17−19

hCtr1 is the main gatekeeper of copper ion uptake into the cells. It binds to copper in the oxidation state of Cu(II). The oxidized Cu(II) form is then reduced to Cu(I) via a mechanism that is not yet fully understood. After the hCtr1 translocated Cu(I) into the cell, specific Cu(I) chaperones deliver it to the dedicated cellular pathways. The first cryogenic electron microscopy (cryo-EM) structure was reported by Unger and co-workers20,21 with a 6 Å resolution revealing the protein to be a trimer containing 50 amino acids in the extracellular N-terminal domain, three transmembrane helices, and a short intracellular C-terminal domain of 15 amino acids (Figure 1). Recently, a 3 Å-resolution crystal structure of hCtr122 without the extracellular domain was reported. The extracellular hCtr1 domain is characterized by several motifs, including glycosylation sites (N15 and T27),23 histidine (His)-rich sites,24 and methionine (Met) motifs.24,25 The His-rich sites in the extracellular domain of hCtr1, specifically the 1MDHSHH and 22HHH segments (Figure 1), were suggested to serve as Cu(II) binding sites.26−30 The Met motifs, 7MGMSYM and 41MMMPM, play a role in Cu(I) binding after the reduction of Cu(II) to Cu(I).29,31 We recently showed that each extracellular domain of hCtr1 binds two Cu(II) ions per monomer by employing in vitro EPR and UV–vis measurements on the full-length hCtr1 protein.32

Figure 1 Schematic picture of the hCtr1 trimer. The extracellular domain of hCtr1 consists of both Cu(II) and Cu(I) sites. The histidine residues form Cu(II) binding sites, and the methionine residues correspond to Cu(I) binding sites. The spin-labeling approach that involves two histidine residues, Cu(II) ion, and NTA ligand is described on the right side.

This study aims to experimentally obtain structural information about the extracellular domain of the full-length hCtr1 protein in vitro and in situ using distance EPR measurements. Cu(II) is a paramagnetic metal ion, and its binding to the extracellular domain of hCtr1 can be used to follow conformational changes. However, Cu(II) is unstable in the reducing environment of the cell and is readily reduced to Cu(I).16 Therefore, an NTA ligand is used to prevent reduction and ensure that Cu(II) binds to the extracellular domain (Figure 1).

Materials and Methods

Cloning, Expression, and Purification of hCtr1

Constructs for the expression of the wild-type (WT) hCtr1 were prepared by PCR amplification and ligated into a modified pFastBac (pK503–9) vector encoding an N-terminal FLAG tag. Point mutations were PCR-amplified using RF cloning.33 To produce baculovirus for hCtr1 expression, a recombinant bacmid was extracted and transfected into Sf9 cells using Cellfectin II Reagent (Thermo Fisher) according to procedures described in the Bac-to-Bac instruction manual (Invitrogen). Insect cells Sf9 were grown at 27 °C in protein-free ESF 921 insect cell culture media (expression systems) in roller bottles and incubated for 3 days postinfection. The cells were harvested and resuspended in a buffer containing 400 mM NaCl, 10% glycerol, 20 mM HEPES (Sigma-Aldrich), pH 7.4, lysed, and centrifuged at 40,000 rpm for 40 min. The pellet was resuspended in a buffer containing 1.5% Triton X-100, 200 mM NaCl, 10% glycerol, 20 mM HEPES, pH 7.4, and incubated overnight at 4 °C. The suspension was centrifuged again at 40,000 rpm for 40 min. CaCl2 (3 mM) was added to the supernatant, which was loaded onto an anti-FLAG M1 agarose affinity gel (Sigma-Aldrich) column, pre-equilibrated with TBS buffer (150 mM NaCl, 50 mM Tris-HCl, pH 7.4), and incubated overnight at 4 °C. The column was washed with TBS buffer, and after elution with 5 mM EDTA-containing buffer, protein-containing fractions were collected and analyzed by sodium dodecyl sulfate polyacrylamide gel electrophoresis (SDS-PAGE) (14% glycine) and silver staining. 250 μM Cu(II) and Cu(II)-NTA were added to a 130 μM purified hCtr1 solution and incubated overnight at 4 °C.

hCtr1 In Situ and Cell Membrane Fragment Experiments

Sf9 insect cells were cultured at 27 °C in protein-free ESF 921 insect cell culture media (Expression Systems) within four roller bottles (50 mL each) and incubated for 3 days postinfection with baculovirus to achieve hCtr1 expression. The cells were then harvested and counted, yielding 8.5 million cells per milliliter. Subsequently, the cells were centrifuged at 21 °C and 1000 rpm for 5 min. The resulting pellet was resuspended in 30 mL of medium, resulting in a final concentration of 28.3 million cells per milliliter. This suspension was then divided into ten test tubes, each containing 5 mL of medium with intact cells. A Cu(II)-NTA solution was prepared by combining 10 mM Cu(II) with 10 mM NTA and mixing overnight. A 250 μM Cu(II)-NTA solution was added to each tube, which was then incubated overnight at room temperature with shaking. The cells were subsequently washed twice with a fresh medium, and samples were kept with 15% glycerol for in situ cell measurements. Additional samples were lysed and subjected to ultracentrifugation at 40,000 rpm and 4 °C for 45 min.

X-Band CW-EPR Measurements

Cu(II) low-temperature EPR measurements were performed using an E500 Elexsys Bruker spectrometer operating at 9.0–9.5 GHz, equipped with a high-sensitivity continuous wave (CW) resonator. Spectra were recorded at low temperatures (130 ± 5 K) at a microwave power of 20.0 mW, a modulation amplitude of 4.0 G, a time constant of 120 ms, and a receiver gain of 60.0 dB. The samples were measured in a 1.6 mm quartz tube (Wilmad-LabGlass, Vineland, NJ) and placed in a 4.0 mm quartz tube for cooling. CW-EPR simulations were carried out using MATLAB, with the EasySpin toolbox.34

Q-Band Double Electron–Electron Resonance (DEER) Experiments

DEER experiments (π/2(νobs) – τ1−π(νobs) – t′ – π(νpump) – (τ1 + τ2 – t′) – π(νobs) – τ2 – echo) were carried out at 20 ± 1.0 K on a Q-band Elexsys E580 spectrometer (equipped with a 2 mm probe head) and 50 W AmpQ. A two-step phase cycle was employed on the first pulse. The echo was measured as a function of t′, whereas τ2 was kept constant to eliminate relaxation effects. The durations of the observer π/2 and π pulses were 14 and 28 ns, respectively. The duration of the π pump pulse was 28 ns, and the dwell time was 12 ns. τ1 was set to 200 ns. The observer frequency was 33.85 GHz, the pump frequency was 33.74 GHz, and the magnetic field was 11,680 G. The samples were measured in 1.6 mm capillary quartz tubes (Wilmad-LabGlass). The data were analyzed using the DeerAnalysis 2019 program. Tikhonov regularization and DEERNet were also used to analyze the data.35

Results and Discussion

The full-length hCtr1 protein was expressed in insect cells, purified by using the agarose affinity gel, and reconstituted into triton micelles as described previously.32Figure S1 shows an SDS-PAGE comparison of the purified protein and the Western blot of hCtr1 from whole cells in the presence of Cu(II) and Cu(II)-NTA. The expression levels of hCtr1 are similar in the presence of free Cu(II) ions and Cu(II)-NTA.

CW-EPR experiments conducted at low temperatures revealed the amino acid residues involved in direct coordination with the copper ion. We added Cu(II) and Cu(II)-NTA at a 2:1 Cu(II):hCtr1 monomer (where the hCtr1 monomer concentration was about 130 μM). Figure 2 shows the low-temperature (130 ± 5K) CW-EPR spectra of Cu(II) and Cu(II)-NTA bound to purified hCtr1. The EPR spectrum suggests that both Cu(II)-NTA and Cu(II) bind in a similar coordination of 2N2O/1N3O to hCtr1 with g⊥ = 2.06, g|| = 2.28, A⊥ = 20 MHz, and A|| = 470 MHz.200 As mentioned previously, the His-rich sites in the extracellular domain of hCtr1, 1MDHSHH and 22HHH, were found to coordinate Cu(II) ions.26−30 The EPR data propose that Cu(II)-NTA complexes bind to these His-rich sites in an analogous way, which is supported by the high affinity of Cu(II)-NTA to His-based segments.16,36 hCtr1 was expressed in insect cells; subsequently, Cu(II)-NTA was added to the cells, and samples for EPR measurements were prepared. The cells were then lysed, and samples of cell membrane fragments containing hCtr1 bound to Cu(ll)-NTA were prepared for EPR measurements. The EPR spectra suggest that Cu(II)-NTA is bound to hCtr1 in the cells and cell membrane fragments under a slightly different coordination environment with the following parameters: g⊥ = 2.05, g|| = 2.25, A⊥ = 20 MHz, and A|| = 480 MHz, indicating 3N1O/2N2O coordination. A low-temperature CW-EPR experiment was also carried out with Cu(II)-NTA added to cells without overexpression of hCtr1, and no specific binding was detected (Figure S2, Supporting Information). This difference indicates that Cu(II)-NTA binds to the additional nitrogen atom on the extracellular domain of hCtr1 in the cells and cell membrane fragments, revealing a different coordination than compared to the purified hCtr1 reconstituted in triton micelles. These data may suggest that while in triton micelles, a single histidine residue may be sufficient to bind Cu(II)-NTA, for in situ and cell membranes, two histidine residues should be involved in Cu(II)-NTA coordination. Three-pulse electron spin echo envelope modulation (ESEEM) (Figure S3, Supporting Information) supports this assignment, where the ESEEM time-domain signals and FT spectra for Cu(II) and Cu(II)-NTA bound to purified hCtr1 are similar and characterized by coordination to 14N nuclei;201 however, the spectra of Cu(II)-NTA in membrane fragments are slightly different and suggest a more pronounced 14N time-domain modulation. As expected, in situ, the signal-to-noise ratio was lower than that in the cell membrane fragments. In both cases, in situ and membrane fragments, clear 14N time-domain modulations were observed.

Figure 2 CW-EPR spectra (acquired at 130 K) for 2:1 Cu(II), Cu(II)-NTA to 130 μM hCtr1 monomer in HEPES buffer, pH 7.4, as well as 250 μM Cu(II)-NTA added to cells with expressed hCtr1 and after lysis. The EPR spectra for purified hCtr1 (in vitro) and cell membrane fragments and in situ are shown as solid lines, and the simulated data are presented as dashed lines. The red and blue dashed lines are drawn to guide the eyes.

Next, Q-band pulsed EPR distance measurements (double electron–electron resonance, DEER) were run on Cu(II) and Cu(II)-NTA bound to purified hCtr1 (Figure 3). The time-domain data and the corresponding distance distribution functions did not reveal any significant differences between the two samples, where bimodal distribution appeared between 1.5 and 3.5 nm. Moreover, additional long-distance contribution between 5.0 and 6.0 nm appeared, which was more pronounced for the Cu(II)-NTA than compared to Cu(II) ions. Overall, based on the CW-EPR and DEER measurements, it seemed that Cu(II)-NTA binds in a similar way to hCtr1 as Cu(II) ions. Therefore, Cu(II)-NTA can be used as a probe for monitoring conformational changes in the extracellular domain of hCtr1.

Figure 3 Q-band DEER measurements on purified hCtr1 (in vitro). Top figures show the presence of free Cu(II), and bottom figures show the presence of Cu(II)-NTA. (A) Time-domain DEER raw data (black) and the background function (red). (B) DEER time-domain data after background correction and the corresponding fit. (C) Corresponding distance distributions. The data were analyzed using the DeerAnalysis program using Tikhonov regularization, where the regularization parameter was 10–20 (solid black lines) and using DEERNet (dashed black lines). Distance distribution validation considered white noise, background start, and dimensionality. The color bar indicates reliability ranges (green: shape reliable; yellow: mean and width reliable; orange: mean reliable; pink: no quantification possible). The data were acquired at 20 K for 2:1 Cu(II), Cu(II)-NTA to 130 μM hCtr1 monomer in HEPES buffer, pH 7.4. 20% glycerol was added.

Subsequently, Q-band DEER measurements were performed on Cu(II)-NTA bound to hCtr1 in situ and cell membrane fragments. First, Cu(II)-NTA was added to the cells at a concentration of 250 μM, which is equivalent to twice the amount of purified hCtr1. After addition of Cu(II)-NTA, the cells were washed and split into two samples: one for in situ EPR measurements of hCtr1 in insect cells and the other one for hCtr1 in cell membrane fragments. Figure 4 shows the DEER time-domain signals and the corresponding distance distribution functions. Interestingly, the small distance distribution functions, below 3.5 nm, that were detected in the purified hCtr1 sample were not observed in situ and cell membrane fragments. In the cell membrane fragments, the DEER data suggested two distance distributions, one around 4.0 nm and another one with a higher population around 5.0 nm, and in situ distributions between 4.6 and 5.8 nm appear. Moreover, the modulation depth, which indicates the number of paramagnetic probes bound to a single protein, is similar to the purified hCtr1 protein and cell membrane fragments (∼1–2%). However, within the cells, the number is much lower (∼0.1%), suggesting that less Cu(II)-NTA molecules are bound to the extracellular domain of the hCtr1 trimer. It is important to note that despite that Cu(II) distance measurements are characterized by low modulation depth, owing to the large spectral width of Cu(II), the use of this spin-labeling methodology allows comparable high-resolution DEER data, owing to the orthogonal labeling, which reduces the flexibility of spin labeling and allows narrow distance distribution functions.37,38

Figure 4 Q-band DEER measurements on Cu(II)-NTA bound to purified hCtr1 and hCtr1 in cell membrane fragments and in situ in insect cells. (A) Time-domain DEER raw data (black) and the background function (red). (B) DEER time-domain data after background correction and the corresponding fit. (C) Corresponding distance distributions. The data were analyzed using the DeerAnalysis program using Tikhonov regularization, where the regularization parameter was 10–20 (solid black lines) and using DEERNet (dashed black lines). Distance distribution validation considered white noise, background start, and dimensionality. The color bar indicates reliability ranges (green: shape reliable; yellow: mean and width reliable; orange: mean reliable; pink: no quantification possible). The data were acquired at 20 K where 250 μM Cu(II)-NTA was added to the cells.

The field sweep EPR spectra (Figure S4, Supporting Information) of purified hCtr1 in the presence of Cu(II) and Cu(II)-NTA were found to be broader than compared to those in the presence of Cu(II)-NTA in cells and cell membrane fragments. Broadening in the EPR field sweep spectrum can originate from paramagnetic centers that are too close, which was also supported by the DEER data. Two-pulse decays (Figure S5, Supporting Information) suggested that the relaxation time of Cu(II)-NTA bound to purified hCtr1 is the shortest, while the relaxation time of Cu(II)-NTA in the cell is the longest. The relaxation time is also dependent on the number of paramagnetic centers bound to a single protein, and therefore, DEER data with higher modulation depth, and a higher number of paramagnetic centers per protein, will be characterized by a shorter relaxation time.

The DEER data derived from in situ and cell membrane fragments suggested that extracellular chains of hCtr1 are further apart than the structure of the purified protein. The extracellular domain is not resolved experimentally; nevertheless, computational prediction of membrane or lipid association revealed a membrane-associated region within the N-terminal end of the protein.39−42 Residues from Ser38 to Phe51 are predicted to be membrane-associated. Interestingly, this sequence from Ser38 to Phe51 also includes five Met residues, which comprise the Cu(I) binding site. Our data suggest that in the native environment, this motif is in proximity with the functional group of the phospholipids and associated with the membrane, resulting in a more distant organization of the three disordered N-termini. Conversely, once the protein is reconstituted in micelles, this anchoring is disrupted and leads to the observed close configuration of the three extracellular domains of hCtr1. Membrane association may be crucial for proper functionality of the protein because these hydrophobic domains can either interact with each other and collapse or are solubilized by micelles which can distort and dislocate the Cu(I) binding site. Recently, EPR spectroscopy showed, by employing various spin-labeling methodologies, that extracellular and intracellular domains of membrane proteins behave differently in reconstituted membrane systems compared to native cell membranes,17,18,43,44 which ultimately influences the gating mechanisms and functionality of membrane proteins.

This study also showed that similar EPR data for a protein in cell membrane fragments and insect cells were obtained. The advantage of measuring native cell membrane fragments compared to whole cells is that the signal-to-noise ratio is improved due to a larger number of paramagnetic centers bound to a single protein.

Conclusions

Following conformational changes in the extracellular domains of membrane proteins is beyond trivial. These domains are disordered, which challenges conventional biophysical tools such as NMR, X-ray crystallography, and electron microscopy. Herein, we apply the new in-cell spin-labeling methodology and EPR measurements to obtain structural information on the extracellular domain of hCtr1. EPR distance measurements were carried out on the purified protein reconstituted in micelles and were compared to that of the protein in membrane fragments and insect cells. We showed that comparable structural data are obtained for the protein in both membrane fragments and intact whole cells, which are both different if compared to the data obtained from the purified protein reconstituted in micelles. These data highlight the importance of gaining structural information on proteins in their native environment.

Supporting Information Available

The Supporting Information is available free of charge at https://pubs.acs.org/doi/10.1021/acs.jpcb.4c03676.Additional control experiments including Western blot, CW-EPR experiments, Q-band 3P-ESEEM experiments, and Q-band two-pulse experiments (PDF)

Supplementary Material

jp4c03676_si_001.pdf

Author Contributions

† S.M. and S.P. have equal contribution.

The authors declare no competing financial interest.

Acknowledgments

S.R. acknowledges the support of the ISF 212/22 grant.
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