To produce functional Hb (haemoglobin), nascent α-globin (αo) and β-globin (βo) chains must each bind a single haem molecule (to form αh and βh) and interact together to form heterodimers. The precise sequence of binding events is unknown, and it has been suggested that additional factors might enhance the efficiency of Hb folding. AHSP (α-haemoglobin-stabilizing protein) has been shown previously to bind αh and regulate redox activity of the haem iron. In the present study, we used a combination of classical and dynamic light scattering and NMR spectroscopy to demonstrate that AHSP forms a heterodimeric complex with αo that inhibits αo aggregation and promotes αo folding in the absence of haem. These findings indicate that AHSP may function as an αo-specific chaperone, and suggest an important role for αo in guiding Hb assembly by stabilizing βo and inhibiting off-pathway self-association of βh.
- α-haemoglobin-stabilizing protein (AHSP)
Hb (haemoglobin) is one of the best studied of all proteins, and a system that has established many paradigms in the relationship between protein structure and function . However, an area that remains comparatively poorly understood is Hb folding and assembly . A specific protein chaperone that may assist Hb folding has been proposed , but the mechanism has not been extensively explored.
Adult human haemoglobin (Hb A) is a tetramer of two α-haemoglobin (αh) and two β-haemoglobin (βh) subunits . The apo-globin chains (αo and βo) lack the haem prosthetic group and precipitate readily, indicating that haem plays a key role in stabilizing the αh and βh structures . The haem-bound chains also require stabilization by binding to each other, as is evident by increased rates of precipitation , elevated haem/iron release  and accelerated autoxidation  of the free chains compared with Hb A. The αh and βh subunits form a strong αhβh heterodimer, which exists in equilibrium with the αh2βh2 tetramer . There is evidence that αhβh assembly proceeds through semi-Hb intermediates, αoβh or αhβo [10,11], which may serve to increase the haem-affinity of apo-subunits . These species confer enhanced stability and possess many functional properties of the intact Hb A .
Although Hb can be reconstituted from apo-globins and haem in vitro, it is possible that other factors are involved in establishing an ordered assembly pathway in vivo to ensure maximum efficiency by, for example, ensuring that haem becomes inserted in the correct orientation . Previously, we obtained genetic and biochemical evidence that AHSP (α-haemoglobin-stabilizing protein) may have a role in Hb assembly . AHSP has been proposed to sequester the free αh that is produced during normal Hb synthesis, and to inhibit its ability to cause oxidative damage [14–17]. This mechanism permits a slight overproduction of αh in order to competitively inhibit the formation of deleterious βh4 tetramers . More recently, we have found that knocking out AHSP expression in both α- or β-thalassaemia mice leads to an exacerbated anaemia [3,15]. This result suggests that AHSP plays an active role in the process of Hb chain assembly, irrespective of the final Hb chain balance. In the present study, we characterize the effects of AHSP binding on αo structure and aggregation state, and compare these with the effects of βh/βo binding.
Protein sample preparation
AHSP, Hb A, αh and βh were produced as described previously . AHSP obtained from this procedure contained the full-length native sequence (102 residues) with an additional Gly-Ser dipeptide at the N-terminus. Hb A, αh and βh were obtained from human blood. To generate αo and βo, purified Hb A was applied to a C4 reversed-phase HPLC column in 35% acetonitrile, 55% water and 0.1% trifluoroacetic acid, and developed with a 35–55% acetonitrile gradient applied over 30 min, similar to the method of Masala et al. . The elution was monitored at 280 and 400 nm and three peaks were eluted in the order: haem, βo and αo. Alternatively, apo-globins were prepared from de-salted αh and βh subunits by acid/acetone extraction, in which the haem is solubilized and the protein fraction is precipitated, collected, washed and dialysed into water . Fractions were freeze-dried and stored at −80 °C before use. Haem contamination was determined by UV–visible absorption spectroscopy in 6 M guanidinium chloride and 10 mM Tris/HCl (pH 8.0); haem concentration was calculated using an absorption coefficient at 390 nm (∊390) of 37800 M−1·cm−1  and polypeptide concentration was obtained using ∊280 values calculated from the amino acid sequence (neglecting the small contribution from residual haem). Both methods yielded a typical haem contamination less that 1% (mol/mol).
To produce 15N-labelled αo for NMR we used the Hb A expression plasmid pHE7, kindly provided by Dr Chien Ho (Carnegie Mellon University, Pittsburgh, PA, U.S.A.), and purified Hb A according to established protocols [22,23]. This method yields Hb A subunits with native N-termini. The 15N-labelled Hb A was purified subject to the same procedures as native Hb A  to obtain αo.
Spectra were collected on a Jasco J-720 spectropolarimeter, using a 1-mm-pathlength cell, and the temperature was controlled using a water-jacketed cell holder and calibrated by means of a thermocouple placed inside the CD cell. CSA (camphor-10-sulfonic acid) was used as a calibration standard. A 1 mg·ml−1 solution of CSA was prepared using an absorption coefficient of 34.6 M−1·cm−1 . Using a 0.1 cm path of CSA standard, the raw ellipticity at 290.5 nm was adjusted to be 33.5 , after correction for the unobstructed beam. The total absorbance was monitored to ensure adequate signal strength across the spectrum (HT < 600). Final spectra were the sum of a minimum of three scans. For the thermal denaturation measurements, the ellipticity at 222 nm was monitored over the temperature range 15–88 °C and a temperature gradient of 1 °C·min−1.
DLS (dynamic light scattering)
DLS measurements were made in 20 mM sodium phosphate buffer (pH 6.3), using a DynaPro™ instrument (Protein Solutions). The time-dependent fluctuation in scattering intensity was fitted to one or more intensity autocorrelation functions to extract the corresponding translational diffusion coefficients (D) using the manufacturer's software. The Stokes radius (RS; the radius of the corresponding hard sphere with the same diffusion coefficient) was calculated according to:
The viscosity of the solution (η) was taken to be 1.005 × 10−2 poise at 20 °C and temperature corrected using the manufacturer's software. Molecular mass (M) was estimated from RS according to: where NA is Avogadro's constant, ῡ is the partial specific volume (assumed to be 0.73 cm3·g−1), and δ is a hydration factor (assumed to be 0.3 g of water per 1 g of protein).
MALS (multi-angle light scattering)
Samples were applied to a Superose 12 column (GE Healthcare) and eluting protein fractions were monitored by inline MALS and RI (refractive index) measurements. MALS intensity was measured at three angles (41.5 °, 90.0 ° and 138.5 °) using a mini-DAWN detector (Wyatt Technology) equipped with a 690 nm laser. Voltages for the three detectors are normalized using BSA monomer (Sigma) as a standard isotropic scatterer. Absolute MALS intensity was calibrated against a standard sample of HPLC-grade toluene. Protein concentrations are obtained from inline RI measurements (Optilab differential refractometer; Wyatt Technology), calibrated against salt solution standards. The RI increment with respect to mass concentration (dn/dc) was taken to be 0.19 ml·g−1 for all proteins/complexes. Weight-average molecular mass (Mw) of the solute was calculated for every 5 μl volume across each eluting peak (a single data ‘slice’) using the Rayleigh–Debye–Gans scattering model for a dilute polymer solution, as implemented by the ASTRA software (Wyatt Technology).
All NMR measurements were conducted on Bruker Avance 600 and 800 MHz spectrometers equipped with 5 mm triple-resonance (1H, 13C, 15N) cryoprobes. Samples were prepared at 0.2–0.4 mM protein concentration in water and adjusted with 10 mM sodium phosphate (pH 6.3) (95% 1H2O, 5% 2H2O; pH values were uncorrected for the presence of 2H2O) with 40 μM DSS (5,5-dimethylsilapentane sulfonate) included as a 0 p.p.m. 1H chemical-shift reference. The 15N chemical-shift scale was referenced indirectly from the DSS. Backbone resonances for αh in complex with AHSP were obtained from HNCA, CBCA(CO)NH, HNCACB and 15N NOESY-HSQC (heteronuclear single-quantum coherence) experiments.
αo preparations adopt a variable α-helical content
To investigate the structure of αo, haem was extracted from purified αh in ice-cold acid/acetone , or by reversed-phase HPLC purification of Hb A . Both methods achieved haem contamination levels of <1% (Figure 1A). At pH 3.2, the far-UV CD spectrum of αo indicated a largely unfolded conformation. Upon raising the pH to 6.3, a spectrum characteristic of α-helical secondary structure was obtained (Figure 1B), consistent with numerous previous studies [5,19,21,26,27]. At 8 °C, CD spectra indicate that αo retains a significant fraction of the native secondary structure compared with αh (Figure 1C, continuous and dashed lines respectively). Raising the temperature to 30 °C induced a significant reduction in α-helical content of αo and an apparent increase in unfolded species, as was evident by a reduction in ellipticity at 192 and 222 nm and a blue shift in the minimum below 208 nm. By comparison, the CD spectra of αh and AHSP showed little variation within this temperature range (results not shown). Unusually, the CD spectrum of αo obtained from different preparations showed significant variation in mean residue ellipticity at 222 nm, and in the 208/222 nm ellipticity ratio. These observations suggest that αo samples readily undergo an irreversible change during purification and storage.
αo contains regions of well-ordered tertiary structure
To investigate the tertiary structure of αo, we produced 15N-labelled Hb A in an Escherichia coli expression system  and purified the 15N-labelled αo subunits for NMR studies. As shown in Figure 1(D), the 1H,15N-HSQC spectrum of αo at pH 3.2 is indicative of an unfolded state, displaying sharp NMR signals with most HN resonances clustered in the range 8–8.7 p.p.m. The number and position of signals is as expected for the 141-residue sequence. Upon raising the pH to 6.3, we saw a general increase in chemical-shift dispersion, indicating the appearance of some well-ordered tertiary structure (e.g. Figure 1E, broken box 1). A number of signals from the pH 3.2 spectrum remain (e.g. Figure 1E, broken box 2). Thus the sample appears to contain both folded and unfolded αo polypeptides. The distribution between the folded and unfolded forms was variable between preparations. A comparison with the 15N-HSQC of αh (Figure 1F) does not reveal any perfectly overlapping signals; however, this is not unexpected considering the large chemical shifts that would be induced by the haem group. Owing to sample instability over a wide range of solution conditions, it was not possible to obtain resonance assignments for the folded form of αo. The average 15N linewidths for both folded and unfolded αo conformers at pH 6.3 are similar at ~18 Hz, and are significantly broadened compared with spectra of αh (14.5±1 Hz), which undergoes monomer–dimer equilibrium, and acid-unfolded αo (8.0±0.5 Hz, presumed monomer), suggesting that folded and unfolded αo species are aggregated or undergoing conformational exchange at pH 6.3.
Unfolding and aggregation of αo is inhibited by AHSP
To investigate the influence of AHSP on the stability of αo, we performed CD thermal melts, monitoring ellipticity at 222 nm. AHSP undergoes a completely reversible thermal unfolding process, with a midpoint temperature of unfolding (Tm) of 57.1±1.7 °C (Figure 2A, dotted line, and Table 1). In contrast, free αo showed a complex, non-co-operative and non-reversible denaturation curve (Figure 2A, continuous line). Addition of AHSP conferred a significant stabilization of αo at physiologically relevant temperatures below 40 °C (Figure 2A, dashed line). Indeed, AHSP was almost as effective at stabilizing αo in this assay as haem (Figure 2B, continuous line). By comparison, AHSP confers little thermal stabilization of αh (Figure 2B, dotted–dashed line). Hb A is considerably more stable than any of the other species, presumably reflecting the effect of the very-high-affinity αhβh dimer.
To investigate the basis for αo stabilization, we monitored the temperature-induced formation of soluble aggregates using DLS to determine RS. The RS measured for αo at 10 °C was considerably larger than expected for a folded globular monomer (Table 1), instead being consistent with aggregated or denatured protein; the empirical relationship RS=0.221N0.57 nm, relating RS to protein chain length (N) for a denatured protein  yields RS=3.7 nm for αo, close to the measured value. AHSP had a RS of 1.7±0.1 nm, consistent with the expectation for a monomeric protein of 12 kDa (Table 1). Addition of 1 molar equivalent of AHSP to αo gave rise to a single species with RS=2.6±0.1 nm, consistent with dissociation of αo aggregates and/or compaction of the αo structure. On raising the temperature, aggregates were seen to develop in free αo solutions (Figure 2C, ▲). Addition of AHSP inhibited this aggregation up to ~35 °C (Figure 2C, Δ), beyond which the sample rapidly converted into large (>100 nm) particles, with visible precipitation. Similar results were obtained for an αh–AHSP complex (Table 1 and Figure 2C). Hb A and AHSP gave no indication of aggregation up to the temperature limit of the DLS apparatus (60 °C) (results not shown).
The stabilizing effect of AHSP on αo precipitation was demonstrated further in a turbidity assay. Proteins were incubated at 25 °C and absorbance at 700 nm was monitored as an indirect measure of light scattering by insoluble material. Precipitate formed rapidly in solutions of αo (Figure 2D, △). Addition of AHSP completely abolished αo precipitation over the 80-min time course of the experiment (Figure 2D, Δ).
αo binds AHSP, βh or βo to form dimeric complexes
To characterize the interactions of αo with potential binding partners in Hb assembly, we performed MALS inline with SEC (size-exclusion chromatography). MALS yields the Mw of proteins in solution, independent of shape or frictional properties. Figure 3(A) shows the use of this method to characterize the interaction between AHSP and αh, known to be a dimer from crystallographic studies. The measured Mw of free αo (18 kDa) is higher than expected for a monomer (15.7 kDa), as a result of weak concentration-dependent self-association (, and results not shown). AHSP has Mw=11.8±0.2 kDa (independent of loading concentration), but elutes considerably earlier than αHb from SEC owing to a more elongated shape (greater RS). Mixing of a 1:1.5 molar ratio of AHSP/αh, followed by SEC, yielded a monodisperse complex with Mw=25.3±1.2 kDa, close to the value expected for a 1:1 complex (Figure 3A, continuous line) and a smaller peak corresponding to unbound αh.
At pH 6.3, αo eluted from SEC in a broad asymmetric peak with a broad distribution of particle sizes ranging from ~20 kDa to over 100 kDa (Figure 3B and Table 1), indicative of non-specific self-association. Addition of an equimolar quantity of AHSP caused a dramatic shift to a discrete αo–AHSP dimer with a Mw of 26±0.2 kDa (Figure 3C). Using this approach, we found that αo also forms a dimeric complex with βh and, in so doing, effectively inhibits the formation of βh tetramers (Figure 3D). Even more surprisingly, αo and βo, which are both unstable and aggregated in isolation, interact to form a more stable αoβo dimer that is eluted as a single peak (Figure 3E). These data highlight αo–AHSP, αoβh and αoβo as potentially important intermediates in Hb A assembly.
AHSP and βh stabilize folded conformations of αo
To probe directly how αo structure is influenced by AHSP and βh, we recorded 15N-HSQC spectra of complexes labelled only on the αo subunit. The αo sample used for this experiment gave an NMR spectrum corresponding to the unfolded species (Figure 4A). Addition of 1 molar equivalent of unlabelled βh resulted in a dramatic reduction in signals associated with this unfolded form, and the appearance of a new set of resonances more characteristic of a well-ordered structure, albeit with a significantly smaller number of peaks than expected for the full αo sequence (Figure 4B). Addition of AHSP produced a similar stimulation of αo folding (Figure 4C, main panel, black). In comparison, addition of AHSP to labelled αh yielded a well-dispersed spectrum with the expected number of peaks (Figure 4C, main panel, red). Backbone 15N, 1HN, 13Cα shifts were obtained to 87% completeness for the αh subunit of αh–AHSP using standard triple-resonance methods. With these data in hand, HNCA and 15N NOESY-HSQC spectra were acquired for αo–AHSP, labelled on the αo subunit. Fewer than 80 backbone HN groups were found that gave rise to detectable NOEs (nuclear Overhauser effects). Short-range NOEs and HN, Cα scalar connectivities were observed for αo residues 14–18 and 20–26 in helices A and B, residues 111–116 linking helices G and H and residues 34–36, and 38 in the helix B and C corner. Chemical-shift similarities with the corresponding residues in αh–AHSP indicated that these regions adopt similar conformations in the two structures. Some additional αo residues could be assigned on the basis of distinctive HN, N, Cα chemical shifts and NOE patterns when compared with αh–AHSP spectra. In total, backbone resonances were assigned for 38 αo residues and mapped on to a previously determined crystal structure of αh–AHSP  (Figure 4D, spheres). It is notable that assignments were not obtained in helix F, or for the majority of helices G and H, indicating that these signals are very weak/absent or have chemical shifts too dissimilar to those from αh–AHSP to be identified. The greatest chemical-shift similarities mapped close to the AHSP interface (Figure 4D, blue and yellow spheres) with larger differences closer to the haem pocket (red spheres), as might be expected.
To probe the αo–AHSP interface further, we reversed the labelling strategy and recorded a 15N-HSQC spectrum of labelled AHSP in complex with unlabelled αo (Figure 4C, inset, black) and compared this with a spectrum of AHSP bound to unlabelled αh (red), assigned in a previous study . Overall, the two spectra were very similar, with coinciding peaks for 73 of the 99 non-proline residues in AHSP (Figure 4D, blue shading on AHSP ribbon; the remaining residues are shaded grey). Most of these peaks were within 0.03 p.p.m. combined weighted 1H, 15N chemical shift, with the maximum difference of 0.1 p.p.m. (see Supplementary Figure S1 at http://www.BiochemJ.org/bj/432/bj4320275add.htm). This indicates that each of these resides lies in a similar physiochemical environment in the αo–AHSP and αh–AHSP complexes. The similarities extended to the subunit interface (see Supplementary Figure S1), indicating that the complementary surfaces of αo/αh, comprising the G and H helices and helix B and C linker, adopt similar structures in each complex.
The structure of αo
Currently, the Hb folding process is not well characterized and, in particular, the potential role of chaperones is not well established. In the present study, we have used a range of biophysical methods to demonstrate that AHSP can promote folding and inhibit aggregation of αo, suggesting key roles for AHSP and αo in regulating Hb assembly.
NMR data indicate the presence of variable proportions of ‘folded’ and ‘unfolded’ forms of αo in typical preparations. Even samples displaying substantially unfolded NMR spectra at pH 6.3 retained α-helical CD signals, suggesting that α-helical conformations persist in αo aggregates not visible by NMR.
Addition of AHSP leads to dissociation of soluble αo aggregates, formation of a stable αo–AHSP dimer and a dramatic shift from unfolded to partially folded forms of αo. When in complex with AHSP, regions of the A, B, C and E helices of αo adopt a stable state that at least partially resembles the native structure on the basis of chemical shifts. In general, assigned residues are surface-exposed, suggesting that buried residues may exhibit weak/absent NMR signals as a result of exchange between alternative packing arrangements. The F helix of αo is particularly notable for the absence of any assigned resonances. The significant degree of α-helical character observed by CD for αo and αo–AHSP, coupled with the large number of broad/absent NMR signals in the ‘folded’ spectra, suggests that regions of αo may sample an ensemble of α-helical conformations on the chemical shift timescale (μs). The G and H helices are presumed to be well-folded in the αo–AHSP complex on the basis of AHSP-binding data, and thus the absence of clear NMR signals in this case may reflect exchange between free and bound αo in addition to internal conformational dynamics.
Thus AHSP can stabilize αo in solution while still retaining a large degree of conformational flexibility in the αo subunit. An attractive possibility is that this serves a functional role by allowing the bulky haem group to access its binding pocket. Such a mechanism has been proposed previously for apo-myoglobin, which retains a near-native fold, except for the F helix, which is disordered until after haem binding . It remains to be determined whether AHSP favourably influences the haem-binding kinetics of αo or improves selectivity for the correct haem orientation. In general, globins traverse very different folding pathways, despite high degrees of similarity in the final protein folds , hence further study of αh and βh folding is required to understand Hb assembly.
The HSQC spectra of αo in complex with AHSP or βh, or in the free folded form, share a number of similar (but not identical) peak patterns, such as peaks arising from Gly15, Gly18, Gly22, Gly25, Gly57 and Thr41 (see Supplementary Figure S2 at http://www.BiochemJ.org/bj/432/bj4320275add.htm). The suggestion is that AHSP and βh stabilize a pre-existing conformation of αo. Interestingly, peptides corresponding to residues 1–86 of αo have been found to be capable of haem binding , supporting the view that the N-terminal region of αo can fold independently of the rest of the protein.
The importance of αo in Hb assembly
Our study of αo provides some insights into potential pathways of Hb assembly. Assembly is thought to proceed through a series of partially haem-saturated intermediates including semi-Hb species, αhβo and αoβh, which exist as dimers in solution [12,34]. Semi-Hb may form through the interaction of αh and βh chains with nascent βo and αo chains, or by incorporation of haem into an αoβo species . However, although dimeric αoβo can be produced in vitro by removal of haem from tetrameric Hb A [5,35,36], purified αo and βo chains were found not to recombine . The role of αoβo as a significant species in Hb assembly is still a matter of debate [2,36]. In our hands, purified αo and βo, each unstable in isolation, do combine spontaneously to form an αoβo dimer that stabilizes both partners, forming what appears to be a discrete 1:1 dimer. In addition, we provide direct evidence that free αo outcompetes βh oligomerization to yield a dimeric semi-Hb αoβh. These data support previously underappreciated roles for αo in Hb assembly, including inhibiting the off-pathway self-association of βh and βo.
Owing to the general instability of free Hb subunits, the efficiency of Hb assembly may be enhanced by the action of specific chaperones . Defects in Hb A production in mouse knockout models [3,14,15] and the finding that Ahsp gene expression is up-regulated under iron-limiting conditions , when free apo-globin chains accumulate, suggest that AHSP plays an important role in Hb A production. The findings of the present study that AHSP stabilizes αo in a partially folded conformation and dramatically increases its resistance to thermal induced denaturation/aggregation provide biochemical support for a role of AHSP as an αo chaperone. We propose that stabilization of αo by AHSP facilitates the formation of αoβo as an important intermediate in HbA assembly, and that αo–AHSP and αoβo complexes might constitute significant routes of entry for haem.
Kaavya Krishna Kumar, Claire Dickson and David Gell performed experiments. Mitchell Weiss, Joel Mackay and David Gell provided the experimental design and wrote the paper.
This work was supported by grants from the Australian Research Council and the National Health and Medical Research Council awarded to D.G.
We thank Ann Kwan and Bill Bubb for expert maintenance of the 600-MHz and 800-MHz NMR spectrometers at the University of Sydney.
Abbreviations: αo, apo-α-globin; αh, α-haemoglobin; βo, apo-β-globin; βh, β-haemoglobin; AHSP, α-haemoglobin-stabilizing protein; CSA, camphor-10-sulfonic acid; DLS, dynamic light scattering; DSS, 5,5-dimethylsilapentane sulfonate; Hb, haemoglobin; Hb A, adult haemoglobin; HSQC, heteronuclear single-quantum coherence; MALS, multi-angle light scattering; NOE, nuclear Overhauser effect; RI, refractive index; SEC, size-exclusion chromatography
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