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Article

Conformational Dynamics of Phytoglobin BvPgb1.2 from Beta vulgaris ssp. vulgaris

by
Simon Christensen
1,*,
Olof Stenström
2,
Mikael Akke
2 and
Leif Bülow
1,*
1
Division of Pure and Applied Biochemistry, Center for Applied Life Science, Department of Chemistry, Lund University, 22100 Lund, Sweden
2
Division of Biophysical Chemistry, Center for Molecular Protein Science, Department of Chemistry, Lund University, 22100 Lund, Sweden
*
Authors to whom correspondence should be addressed.
Int. J. Mol. Sci. 2023, 24(4), 3973; https://doi.org/10.3390/ijms24043973
Submission received: 30 November 2022 / Revised: 9 February 2023 / Accepted: 12 February 2023 / Published: 16 February 2023
(This article belongs to the Section Molecular Biophysics)
Browse Figures
Versions Notes

Abstract

:
Plant hemoglobins, often referred to as phytoglobins, play important roles in abiotic stress tolerance. Several essential small physiological metabolites can be bound to these heme proteins. In addition, phytoglobins can catalyze a range of different oxidative reactions in vivo. These proteins are often oligomeric, but the degree and relevance of subunit interactions are largely unknown. In this study, we delineate which residues are involved in dimer formation of a sugar beet phytoglobin type 1.2 (BvPgb1.2) using NMR relaxation experiments. E. coli cells harboring a phytoglobin expression vector were cultivated in isotope-labeled (2H, 13C and 15N) M9 medium. The triple-labeled protein was purified to homogeneity using two chromatographic steps. Two forms of BvPgb1.2 were examined, the oxy-form and the more stable cyanide-form. Using three-dimensional triple-resonance NMR experiments, sequence-specific assignments for CN-bound BvPgb1.2 were achieved for 137 backbone amide cross-peaks in the 1H-15N TROSY spectrum, which amounts to 83% of the total number of 165 expected cross-peaks. A large proportion of the non-assigned residues are located in α-helixes G and H, which are proposed to be involved in protein dimerization. Such knowledge around dimer formation will be instrumental for developing a better understanding of phytoglobins’ roles in planta.

1. Introduction

Hemoglobins (Hb) are essential for almost all forms of life. These heme-equipped proteins are not limited to mammals, but are found in a wide range of living organisms, spanning from vertebrates and plants to protozoans and bacteria [1]. Several metabolic roles have been attributed to Hbs in addition to binding a variety of compounds including oxygen, carbon monoxide (CO), nitric oxide (NO), hydrogen sulfide (H2S), hydroxylamine (NH2OH), and several other small organic molecules [2,3]. Hbs are also able to catalyze a range of oxidative reactions thereby suggesting that these proteins show a large diversity of physiological functions [4,5,6].
Particularly plant Hbs have attracted substantial scientific attention during the last decade. However, already in the 1930s, the first plant Hb was discovered, a symbiotic Hb from soybean [7]. Since then, different classes of plant Hbs, also called phytoglobins (Pgb) [8], have been identified. Several physiological functions have been proposed for these proteins, including a roles in adaptation and survival during hypoxia [9,10,11], metabolism of nitrogen [12,13], and hormone/signaling pathways [13,14,15].
Hbs are composed of a heme b prosthetic group inserted within an α-helical protein structure composed of eight helices, A–H. The binding between the heme group and the globin chains differ. Class I Pgb proteins have a low hexa-coordination equilibrium constant (KH), which describes the binding constant of the distal histidine and hence the equilibrium of the penta- and hexa-coordinated states [16]. The low KH results in high affinity for oxygen, which is optimally designed for oxygen-dependent nitric oxide scavenging in low oxygen-containing environments [17,18]. Furthermore, the slow oxygen dissociation rate makes class I Pgs unlikely to function as oxygen transporters. Instead, Class I Pgb play an important role in electron transport due to their high oxygen affinity and redox potential, which serve to maintain the energy status of the cell during oxygen deprivation [7,18]. However, this reasoning is not universally accepted. Instead, recent research of Pgbs from sugar beet suggests that even if the Hbs have intrinsic properties to catalyze certain oxidative reactions, the location and concentration of ligands, such as oxygen, NO, and nitrite will ultimately determine their activity [13].
Over the years, the biennial sugar beet (Beta vulgaris spp. vulgaris) has gathered substantial interest due to its socioeconomical importance for both sugar and energy production. In this caryophyllidae, three different Pgbs have been found (BvPgb1.1, BvPgb1.2, and BvPgb2). Their gene expression pattern differs depending on, e.g., the growth phase and photoperiods of the plant [3]. In a study by Leiva-Eriksson et al. [13], the characteristics of these relatively recently discovered Pgbs were determined and compared to existing data for Pgbs from both eudicots and monocots. Here, it was found that BvPgb1.2 (Pgb class 1) has a lower equilibrium binding constant for oxygen (KO2) compared to the common model Hb protein in Arabidopsis, AtHb1, and to monocotic Pgbs from barley, rice, and maize [13]. Furthermore, BvPgb1.2 also exhibited a lower hexa-coordination constant (KH) in comparison with proteins found in Arabidopsis thaliana and the mentioned monocots.
In addition, the class II Pgb from sugar beet (BvPgb2) was recently heterologous expressed in yeast, A. thaliana and tomato (Solanum lycopersicum). In the case of A. thaliana, such overexpression conferred improved drought and osmotic stress tolerance. The enhanced levels of BvPgb2 in tomato both increased iron levels in leaves and made the plants more resistant to drought-induced withering [19]. Since tomato is the most consumed horticultural crop worldwide, there is a strong driving force to lower its high water and irrigation requirements and make the crop more resilient to external abiotic stress factors [19]. In addition, when this protein was overexpressed in field cress (Lepidium campestre), an increased oil content was observed [20]. These results on BvPgbs have provided important information on the diversity of Pgbs. The metabolic roles of these proteins and other Hbs in crops have indicated their potential to improve agricultural performances [21,22]. Furthermore, food quality in terms of enhanced iron levels may promote the use of these proteins as additives in various products.
Due to growing importance of Pgbs it is essential to further delineate possible structure-function relationships to explain their physiological functions. We recently solved the three-dimensional structure of BvPg1.2 [23] in its monomeric form. Since this Pgb is largely present in a dimeric form, it is essential to gain more knowledge on the subunit interactions present in the protein. Nuclear magnetic resonance (NMR) spectroscopy is a powerful tool to examine molecular structure, dynamics and interactions [24]. In the field of protein science, NMR spectroscopy hence provides additional information on dynamics that complements structural information obtained by X-ray crystallography. NMR relaxation methods probe internal motions within the molecule that are often essential for function [25,26]. One common obstacle in protein NMR is the size limitation caused by spectral overlap among signals and line-broadening, both of which increase with molecular size. However, by labeling the protein with stable isotopes (15N and 13C) the overlap problem can be mitigated, and labeling with 2H significantly reduces line-broadening, enabling NMR studies of proteins up to 40–50 kDa [27,28].
In this study, we report on the conformational dynamics of BvPgb1.2 in its cyanide-bound form, based on backbone 15N longitudinal (R1) and transverse (R2) NMR relaxation rate measurements. In addition, we compare the spectral characteristics of the oxy- and cyanide-bound forms. The main objective was to expand the current knowledge regarding dimer–dimer interactions and identifying dynamic motions/flexible regions in this class of Pgbs. The results clearly indicate which protein regions are involved in dimer formation and highlighting dynamic movements in BvPgb1.2. Since BvPgbs can interact with a range of other biomolecules in vivo, including nucleic acids, lipids, and other proteins, the main objective was to further dissect the subunit interaction areas which most likely are involved in such interactions.

2. Results and Discussion

We assigned the NMR spectrum for CN-bound BvPgb1.2 using three-dimensional triple-resonance experiments (see Section 3). We achieved sequence-specific assignments for the cross-peaks from 137 backbone amides in the 1H-15N TROSY spectrum, Figure 1, which amounts to 83% of the total number of 165 expected cross-peaks (171 residues minus 6 prolines). For an additional 13 residues, we could assign one or several of the 13Cα, 13Cβ, and 13C’ (carbonyl) chemical shifts, while 15 non-proline residues remain unassigned. There are several unassigned peaks in the 1H-15N TROSY spectrum indicating that there are one or a few broadened peaks separating the unassigned from assigned peaks.
By comparison, in the 1H-15N TROSY spectrum of oxygen-bound BvPgb1.2, we could assign only 48 out of the 165 expected cross-peaks from backbone amide groups. The lack of assignment can be explained by the paramagnetic property of the ferrous (FeII) heme, which is in a high-spin state when bound to oxygen [29,30], causing significantly increased relaxation of the nuclear spins. In contrast, CN-bound BvPgb1.2 is in the diamagnetic, low-spin ferric state (FeIII) [31], resulting in slower relaxation and improved NMR spectra. This effect is further illustrated by Figure 2, which shows assigned residues color coded in blue on the structure; the assigned residues are those located furthest away from the heme-group where oxygen is bound [32].
In what follows, we focus our report on the dimerization of CN-bound BvPgb1.2 and its conformational dynamics.

2.1. NMR Spectroscopy Provides Evidence for Dimerization of CN-BvPgb1.2

All residues with missing assignment or partial assignment in CN-bound BvPgb1.2 are listed in Table 1 and mapped onto the structure in Figure 2A. One notable missing assignment is the proximal histidine H112, which coordinates the iron of the heme group together with the distal histidine H77. Assignments are also missing for residues 129–133 located in α-helix G, and for residues 150–156 in the adjacent helix H. The preceding residues 144–149 appear significantly broadened or have a shoulder, suggesting that they are present in two conformations. Moreover, residues in the N-terminal segment, located next to helix H, also show evidence of populating multiple conformations: V6 shows three peaks in the 1H-15N TROSY spectrum and residues 7–17 all show two peaks, except for G13, which was assigned to a single peak, and I16, which is missing in the spectrum. Residues showing multiple sets of peaks or peak broadening are listed in Table 1 and mapped onto the structure in Figure 2B. Notably, residues with conformational heterogeneity map to a contiguous region of the protein structure, which has previously been implicated in mediating dimerization of plant Hbs. The dimer dissociation constant (Kd) has been determined to 86 μM for rice Hb1 in the ferric state [33] (63.4% sequence identity compared to BvPgb1.2). Based on this value, we estimate that the fraction of monomeric protein in the NMR sample of CN-bound BvPgb1.2 is around 20% of the total protein concentration. Furthermore, the dimer dissociation constants for phytoglobins from Parasponia andersonii, Trema tomentosa, and Arabidopsis thaliana (72.2, 74.1, and 74.4% sequence identity, respectively) have been determined and shown to be mutually similar with Kd ~1 μM [34,35,36]. Using this Kd value, one would predict that there is only ~3% monomer in the BvPgb1.2 NMR sample.
A large proportion of the non-assigned residues is situated in α-helixes G and H, which in other Hbs are involved in dimerization [33]. A likely explanation for the missing assignments is exchange broadening due to transient dimerization or conformational exchange at the dimer interface. Chemical exchange has been observed previously in the corresponding structural region of other Hbs, and has been explained by a sliding motion involving the two protomers or by slow diffusion of low-molecular weight molecules, such as O2, NO or CO, in ‘tunnels’ forming between helices G and H [37,38]. The latter explanation is unlikely in the present case, given the sample conditions, while the former is a possible explanation for the conformational heterogeneity observed in the NMR spectra. Note that exchange due to transient dimerization or sliding motions are not mutually exclusive. Several NMR studies have also observed exchange in the F-helix [37,39]. It is certainly plausible that the four unassigned residues in helix F of CN-bound BvPgb1.2 might be broadened by conformational exchange. The exchange timescale(s) are not well defined: the observation of multiple peaks (or shoulders) suggests that the underlying exchange is slower than the difference in chemical shift between the alternative states, whereas the exchange broadening beyond detection might suggest a relatively fast process, or that the chemical shift difference in these cases is smaller. As mentioned above, it is possible that there are two distinct processes active: transient dimerization, which might be fast, and local conformational exchange, which might be slower, or vice versa. We address this issue next using 15N spin relaxation methods.

2.2. 15N Relaxation Confirms Dimerization and Identifies Flexible Regions

To characterize the conformational dynamics of CN-bound BvPgb1.2, we measured longitudinal (R1) and transverse (R2) 15N relaxation rate constants that probe motions on the picosecond to nanosecond and millisecond time scales. Out of the 137 assigned residues, 7 residues (N7, S11, Q72, K73, L74, L105, and E123) were excluded from further analysis due to severe spectral overlap. The remaining 130 residues yielded high quality relaxation data (Figure 3A,B), albeit with relatively large error bars for a handful of residues with higher than average values of R2 (Figure 3B), as might be expected due to the increased line-broadening.
We determined the rotational correlation time (τc) from the residue-specific ratios of R2/R1 and the structure of BvPgb1.2 (pdbID: 7ZOS [23]), using the program ROTDIF (Table 2), based on a dataset trimmed by removing all R2/R1 values outside of one standard deviation from the mean [40]. The trimmed data set comprises 104 residues. The best fitting diffusion model is anisotropic with τc = 22.8 ns, an anisotropy of 0.84, and a rhombicity of 0.63. We compared τc determined by ROTDIF with τc calculated using HYDRONMR based on the monomeric structure of BvPgb1.2 (PDB-id: 7ZOS [23]) and the dimeric structure of BvPgb C86A (PDB-id: 7Z1U [23]). HYDRONMR predicts τc values of 11.3 ns and 28.9 ns for the monomeric and dimeric structures, respectively, in agreement with correlation times observed for other Hbs [39,41]. The anisotropy and rhombicity was 0.83 and 0.40 for the monomeric structure and 1.44 and 0.12 for the dimeric. Together, these results suggest that CN-BvPgb1.2 exists primarily in dimeric form in solution at the present concentration, as is also found for other phytoglobins [29,31,42]. This conclusion is also in agreement with the weak intensity of the minor cross-peaks in the NMR spectrum, which indicate that the monomer population must be low, if the system is in slow exchange. Based on results from this and previous work [23], we suggest hydrogen bonding primarily between T53-E120 in opposing dimers but H121-E120 could also interact in this way. Moreover, a hydrophobic cluster between I47, I50, V124 and F127 aids in dimer formation through hydrophobic interactions, even though additional residues with similar properties might facilitate quaternary stabilization as well. All these residues are primarily located in the B/C loop and G helix, in line with the previous hypothesis [36].
The relaxation data show increased flexibility for some 25 residues with low values of R2/R1 (Figure 3C). The N and C termini are both highly flexible. Other positions with low R2/R1 ratios include residues 28, 34–35, 63–66, 99, 101, 104, 119, and 124. For most of these residues, the increased flexibility can be explained by structural features: K34 and N35 are located in the kink region between α-helices A and B; 63–66 are situated between helixes C and E in a region that is lacking electron density in the crystal structure, indicating high flexibility; T99, F100, and G101 reside in the loop region between helixes E and F; S103 and S104 are situated at the N-terminal end of α-helix F. Two residues, S28 and V124, with increased flexibility are more difficult to explain in terms of structure, as they are located in the middle of α-helices A and G, respectively.
Elevated R2/R1 ratios are observed for residues L39, I47, E49, I50, K55, F60, V81, M84, K87, S88, Q91, R93, M108, S110, V111, F122, and W149. Residues L39, I47, E49, and I50 are all located in non-central parts of helix B, while K55 is in helix C, close to the heme group, but not directly interacting with it. F60 is located at the beginning of the loop directly following helix C. V81, M84, K87, S88, Q91, and R93 are all located on the same side of helix E, consecutively separated by one turn of helix and connected by hydrogen bonds. V81 is located one turn of helix away from the heme-coordinating H77. It is possible that the pattern of R2/R1 observed for these residues suggests transient bending of the helix. A similar pattern is observed for residues M108, S110, and V111 in helix F, which also harbors H112, one of the other heme-coordinating residues. Furthermore, residues V81, M84, M108, V111, and F122 have their side chains lining the heme-binding pocket. Taken together, these results might suggest that the heme-binding pocket is undergoing some type of breathing motions, accompanied by transient bending or fraying of the neighboring helices.
In a recent study, we investigated the structural and functional roles of C86 in BvPgb1.2 by comparing the wild-type protein with a C86A mutant [23]. Wild-type BvPgb1.2 crystallized as a monomer, while the C86A mutant (PDB-id:7Z1U [23]) crystallized as a dimer, with only minor structural differences observed between the two variants. The dimeric structure emphasizes the importance electrostatic and hydrophobic interactions between the B/C and G helices in the neighboring subunit [36]. Our results showed that while these two variants have very similar three-dimensional structures, the autoxidation rate is increased for the C86A mutant, which also has a lower peroxidase activity. In addition, Y115 has been suggested to control the redox stability of the iron together with C86 [43,44]. While the aromatic ring of Y115 is directly interacting with the heme, the side chain of C86 is located between helices A and E and is pointing away from the heme towards the surface of the protein. Both these residues have relaxation rates similar to the average value observed for other central residues in helices, indicating that their dynamics do not stand out. However, both residues might still be affected by the dynamics observed in helices E and F, described above: it is likely that bending or fraying of helix E would affect the solvent accessibility of the C86 sulfhydryl group, thereby influencing its oxidation state. Similarly, conformational dynamics of helix F might well modulate the interactions between the Y115 side chain and the heme.

3. Materials and Methods

3.1. Protein Expression and Purification

3.1.1. Construction and Transformation of Expression Constructs

The gene for Beta vulgaris spp. vulgaris Pgb with GenBank Accession KF549981 (BvPgb1.2) [13] was custom synthesized by Epoch Biolabs (Missouri City, TX, USA). Recombination based cloning and transformation were carried out according to manufacturer’s instructions (GatewayTM, Invitrogen, Waltham, MA, USA). The gene contained attB1 and attB2 sites at the 5′ and 3′ ends, respectively. The genes were individually cloned into a pET-DEST42 plasmid. The final expression vector was transformed into BL21-DE3 E. coli cells.

3.1.2. Expression of Triple-Labeled Protein in Minimal Media

The cells containing the expression vectors were grown in conventional M9 minimal media containing 15N- or 13C/15N-labeled components. The adaptation of E.coli BL21-DE3 cells were based on previous work with some modifications [45]. (i) Cells were inoculated in LB medium overnight. (ii) 15 mL non-deuterated M9 medium was inoculated with cells from the overnight culture. The cells were grown ~8 h and centrifuged 5 min at 10,000 rpm, followed by resuspension in fresh M9 media for another 8 h. (iii) Cells were centrifuged as before and suspended in M9 medium containing 70% D2O and grown for ~8 h or until sufficient cell mass was obtained. This step was done twice. (iv) Cells were suspended in M9 medium containing 99.8% D2O in the same way as before. This was done twice. (v) Cells were suspended in M9 medium containing 0.8% w/v 13C-glucose, 2 g/L 15NH4Cl, and 99.8% D2O. This was done three times, where a 15% glycerol stock of adapted cells were frozen and kept. The final cultivation volume was 3 L of triple-labeled M9 medium and shake flasks were inoculated using adapted cells from step v.
Cells containing the expression vectors were grown in triple-labeled minimal medium containing 100 µg/mL carbenicillin at 37 °C and 150 rpm until A600 ≥ 2. Expression of BvPgb1.2 was then induced by adding 0.5 mM IPTG and 0.3 mM δ-aminolevulinic acid. The cells were also briefly bubbled with CO to obtain a stable CO-BvPgb when expressed. After induction, cells were grown for 48 h at 22 °C, 150 rpm, then collected by centrifugation, and finally frozen at −80 °C until used.

3.1.3. Cell Harvesting and Protein Extraction

Cell harvesting and protein extraction for the triple-labeled protein was conducted using previously described methods [13].

3.1.4. Protein Purification

The extracted supernatant was dialyzed twice in 4-L 50 mM Tris-HCl pH 8.5 buffer during 12h before protein purification. Dialyzed lysates were then passed through 1.2 µm filter (Sarstedt, Nümbrecht, Germany). BvPgb1.2 were lastly separated and purified from the lysates through a two-step chromatography process consisting of an initial anion-exchange step followed by a hydrophobic interaction chromatography step. Both steps were performed on an ÄKTA™ Avant (Cytvia Life Science, Singapore) chromatography system. All buffers used for chromatography were pH adjusted and degassed, and all solutions were passed through a 0.45 µm filter before being loaded onto columns. Filtered lysates were first loaded onto a Q-Sepharose FF column (Cytvia Life Science) and eluted with three column volumes (CV) of 50 mM Tris-HCl pH 8.5 and 200 mM NaCl. The eluted BvPgb1.2 fractions were pooled and concentrated using 10 kDa Vivaspin® 20 mL ultrafiltration units (Vivascience, Stonehouse, UK) before adding ammonium sulfate (NH4)2SO4 to a final concentration of 1 M. The fractions were then loaded onto a Butyl-Sepharose HP column (Cytvia Life Science) pre-equilibrated with 50 mM Tris-HCl pH 8.5 buffer and 1.5 M (NH4)2SO4.
BvPgb1.2 were then eluted with a three CV linear gradient ranging from 1 M to 0 M (NH4)2SO4 in 50 mM Tris-HCl pH 8.5 buffer. The final BvPgb1.2 fractions had an absorbance ratio at 412 nm and 280 nm (A412/A280) in the range 2.5–3, corresponding to >98% purity. BvPgb1.2 fractions were lastly concentrated using 10 kDa Vivaspin® 20 mL ultrafiltration units (Vivascience) and stored at −80 °C. When examined by analytical size exclusion chromatography the protein samples were identified as dimers [23].

3.2. Protein Sample Preparation

To prepare the cyanide form of BvPgb1.2, the purified protein was dialyzed in 10 mM potassium ferricyanide and 1 mM potassium cyanide dissolved in 50 mM MOPS for 8 h in 0.5 L solution; this process was performed twice. The cyanide-protein solution was passed through a PD10 column (Cytvia Life Science) to remove excess cyanide and the protein was concentrated using 10 kDa Vivaspin® 20 mL ultrafiltration units (Vivascience). The oxy form of BvPgb1.2 was generated by passing the purified Hb through a PD10 column (Cytvia Life Science) in order to equilibrate with the oxygenated buffer (50 mM MOPS).

3.3. NMR Spectroscopy and Data Analysis

3.3.1. NMR Sample Preparation

NMR samples of CN-BvPgb1.2 contained 1 mM protein in 400 μL 50 mM MOPS pH 7.4 with 8% D2O added for the field-frequency lock system on the NMR spectrometer.

3.3.2. Triple-Resonance NMR Experiments and Data Analysis for Backbone Assignment

Three-dimensional triple-resonance 1H/13C/15N NMR experiments for backbone resonance assignments were carried out at 298 K on a Bruker Avance HDIII 800 MHz spectrometer, equipped with a TCI 800S7 H-C/N-D-03 Z probe. A series of 1H–15N TROSY-based three-dimensional 1H-detected experiments with 2H decoupling were acquired using targeted acquisition [46] and random non-uniform sampling. The experiments included HNCO (25% and 30% sampling density), HN(CO)CA (12% and 13.9% sampling for the CN-bound and oxygen-bound samples, respectively), HNCA (12% and 12.9%), HNCACB (23% and 26%), HN(CO)CACB (24% and 26%), and HN(CA)CO (11% and 13.1%). The data were processed using the compressed sensing IRLS algorithm in MddNMR [47,48]. Residue-specific assignment was carried out using the CcpNmr Analysis software suite [49].

3.3.3. 15N NMR Relaxation Experiments and Data Analysis

1H–15N TROSY-based 15N R1 and R2 relaxation experiments targeting the backbone amides [50] were performed at a static magnetic field strength of 14.1 T and a temperature of 298 K. Spectral widths were 14.4 ppm and 30 ppm, covered by 2048 and 256 points in the 1H and 15N dimensions, respectively. Relaxation decays were recorded using 10 delays with times ranging from 0.020–1.2 s and 0–220 ms for R1 and R2, respectively. Spectra were processed using NMRPipe [51] and a processing protocol including zero filling to twice the number of data points in both dimensions, solvent filter, and a squared cosine apodization. Relaxation rates were calculated using PINT, which can resolve overlapped peaks using line fitting [52]. Peak intensities were evaluated using a weighted sum of Gaussian and Lorentzian lineshape functions. Error estimation of the fitted relaxation rates was performed using the jack-knife procedure.
HydroNMR [53] calculations were performed using version 7c through NMRBox [54] at 14.1 T and with atomic element radius (AER) of 3.1 Å. We determined the τc of the BvPgb1.2 by fitting the rotational diffusion tensor to the 15N R1 and R2 values using the JAVA version of Rotdif 3.1 [55,56]. The 1H–15N bond vector orientations were taken from the structure of BvPgb1.2 to which hydrogens were added using the AddH function in Chimera [57].

4. Conclusions

In this study, we investigated the conformational dynamics in sugar beet hemoglobin BvPgb1.2, the first phytoglobin to be characterized using NMR in an extensive way. A proportion of the non-assigned residues is situated in α-helixes G and H, where tunnel forming between these helices might occur, leading to diffusion of low-molecular weight molecules. Correlation time determination using ROTDIF and HYDRONMR revealed similar correlation times compared to a dimeric phytoglobin, suggesting that BvPgb1.2 exists mainly as a dimer for these conditions. Results from 15N relaxation indicated a dynamic motion of the heme pocket which might occur in bending and fraying of the neighboring helices. However, previously studied residues that might be important for redox environment and electron acceptors around the heme pocket (C86 and Y115) displayed relaxation rates that did not stand out from average values, although it is still possible that the bending of the E and F helices might affect solvent accessibility and modulate interactions for these residues. Taken together, the results revealed important clues into the dynamic motions of these proteins and how these conformations might lead to subunit interactions in phytoglobins.

Author Contributions

S.C. performed protein expression, purification and prepared samples. M.A. and O.S. planned the NMR experiments, O.S. conducted NMR experiments, M.A., O.S. and S.C. interpreted and analyzed the data. S.C., O.S., M.A. and L.B. planned the overall project. All authors have read and agreed to the published version of the manuscript.

Funding

This research was funded by Swedish Research Council for Environment Agricultural Sciences and Spatial Planning, grant number 2020-0185, and the Swedish Research Council, grant number 2019-03996.

Institutional Review Board Statement

Not applicable.

Informed Consent Statement

Not applicable.

Data Availability Statement

Experimental data can be obtained from the corresponding auhors on reasonable request.

Acknowledgments

NMR spectroscopy was carried out in part at the Swedish NMR Center, University of Gothenburg. We thank Cecilia Persson for assistance with the NMR experiments.

Conflicts of Interest

The authors declare that the research was conducted in the absence of any commercial or financial relationships that could be construed as potential conflicts of interest.

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Ijms 24 03973 g001 550
Figure 1. 1H–15N-TROSY spectra with assignments for BvPgb1.2 in complex with cyanide (A,B) and oxygen (C). The spectra were acquired at 298 K and a magnetic field strength of 18.8 T. Panel B is a close-up of the boxed area in panel A.
Figure 1. 1H–15N-TROSY spectra with assignments for BvPgb1.2 in complex with cyanide (A,B) and oxygen (C). The spectra were acquired at 298 K and a magnetic field strength of 18.8 T. Panel B is a close-up of the boxed area in panel A.
Ijms 24 03973 g001
Ijms 24 03973 g002 550
Figure 2. (A) High R2 (blue), high R1 (red), missing (green), and partial (cyan) assignments and (B) residues showing two conformations or line broadening (purple) color coded on the structure of BvPgb1.2.
Figure 2. (A) High R2 (blue), high R1 (red), missing (green), and partial (cyan) assignments and (B) residues showing two conformations or line broadening (purple) color coded on the structure of BvPgb1.2.
Ijms 24 03973 g002
Ijms 24 03973 g003 550
Figure 3. NMR relaxation data R1 (A), R2 (B), and R2/R1 (C) measured at 600 MHz, plotted against residue number. Secondary structure elements along the sequence are shown on top, where alpha helices are represented by purple boxes and loop regions by black lines.
Figure 3. NMR relaxation data R1 (A), R2 (B), and R2/R1 (C) measured at 600 MHz, plotted against residue number. Secondary structure elements along the sequence are shown on top, where alpha helices are represented by purple boxes and loop regions by black lines.
Ijms 24 03973 g003
Table
Table 1. List of residues with partial assignments and missing assignments.
Table 1. List of residues with partial assignments and missing assignments.
Partial assignmentsN5 P9 I16 I47 F48 T53 P67 T85 E102 K114 P138 P143 V157
Missing assignmentsM1 S2 F3 T4 P52 H112 L113 L129 L130 E131 T132 I133 A150 E151 A152 F153 N154 H155 L156 I160 P171
Resides with more than one conformation V6 N7 Y8 A10 S11 D12 G13 T14 V15 F17 V98 E144 M145 K146 N147 A148 W149 R166 L167 S168
Table
Table 2. Properties of diffusion tensor determined by ROTDIF and HYDRONMR.
Table 2. Properties of diffusion tensor determined by ROTDIF and HYDRONMR.
τc (ns)AnisotropyRhombicity
ROTDIF22.80.840.63
Monomer (pdbID: 7ZOS)11.30.830.40
Dimer (pdbID: 7Z1U)28.91.440.12
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Christensen, S.; Stenström, O.; Akke, M.; Bülow, L. Conformational Dynamics of Phytoglobin BvPgb1.2 from Beta vulgaris ssp. vulgaris. Int. J. Mol. Sci. 2023, 24, 3973. https://doi.org/10.3390/ijms24043973

AMA Style

Christensen S, Stenström O, Akke M, Bülow L. Conformational Dynamics of Phytoglobin BvPgb1.2 from Beta vulgaris ssp. vulgaris. International Journal of Molecular Sciences. 2023; 24(4):3973. https://doi.org/10.3390/ijms24043973

Chicago/Turabian Style

Christensen, Simon, Olof Stenström, Mikael Akke, and Leif Bülow. 2023. "Conformational Dynamics of Phytoglobin BvPgb1.2 from Beta vulgaris ssp. vulgaris" International Journal of Molecular Sciences 24, no. 4: 3973. https://doi.org/10.3390/ijms24043973

APA Style

Christensen, S., Stenström, O., Akke, M., & Bülow, L. (2023). Conformational Dynamics of Phytoglobin BvPgb1.2 from Beta vulgaris ssp. vulgaris. International Journal of Molecular Sciences, 24(4), 3973. https://doi.org/10.3390/ijms24043973

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