Structural and Catalytic Effects of Proline Substitution and Surface Loop Deletion in the Extended Active Site of Human Carbonic Anhydrase II

Abstract

The bioengineering of a thermophilic enzyme starting from a mesophilic scaffold has proven to be a significant challenge as several stabilizing elements have been proposed to be the foundation of thermal stability including disulfide bridges, surface loop reduction, ionic pair networks, proline substitutions, and aromatic clusters. This study emphasizes the impact of increasing the rigidity of human carbonic anhydrase II (HCA II) via incorporation of proline residues at positions 170 and 234, which are located in surface loops that are able to accommodate restrictive main-chain conformations without rearrangement of the surrounding peptide backbone. Additionally, the effect of compactness of HCA II was examined by way of deletion of a surface loop (residues 230 through 240), which had been previously identified as a possible source of thermal stability for the hyperthermophilic CA isolated from the bacterium Sulfurihydrogenibium yellowstonense YO3AOP1. Differential scanning calorimetry analysis of these HCA II variants revealed that these structural modifications had a minimum effect on the thermal stability of the enzyme while kinetic studies showed unexpected effects on the catalytic efficiency and proton transfer rates. X-ray crystallographic analysis of these HCA II variants showed the electrostatic potential and configuration of the highly acidic loop (residues 230 and 240) plays an important role in its high catalytic activity. Based on these observations and the literature, a picture is emerging of the various components within the general structural architecture of HCA II that are key to stability. These elements could provide the blueprints for the rational thermal stability engineering of other enzymes.

Introduction

Human carbonic anhydrase II (HCA II) is zinc-containing metalloenzyme that catalyzes the reversible hydration of carbon dioxide (CO₂) into bicarbonate (HCO₃⁻) and a proton (H⁺) via a two-step ping-pong mechanism [1, 2]:

$$\mathrm{C}{\mathrm{O}}_2+\mathrm{E}\mathrm{Z}\mathrm{n}\mathrm{O}{\mathrm{H}}^{-}\leftrightharpoons \mathrm{E}\mathrm{Z}\mathrm{n}\mathrm{H}\mathrm{C}{{\mathrm{O}}_3}^{-}\leftrightharpoons \mathrm{E}\mathrm{Z}\mathrm{n}{\mathrm{H}}_2\mathrm{O}+\mathrm{H}\mathrm{C}{{\mathrm{O}}_3}^{-}$$ (1)

$$\mathrm{E}\mathrm{Z}\mathrm{n}{\mathrm{H}}_2\mathrm{O}+\mathrm{B}\leftrightharpoons \mathrm{E}\mathrm{Z}\mathrm{n}\mathrm{O}{\mathrm{H}}^{-}+\mathrm{B}{\mathrm{H}}^{+}$$ (2)

In the first step of the reaction, in the hydration direction, CO₂ binds in a hydrophobic pocket at the base of the active site, with the carbon positioned for nucleophilic attack by a zinc-bound hydroxide (Zn-OH⁻) leading to the formation of HCO₃⁻. The HCO₃⁻ is then displaced from the zinc and leaves the active site with water taking its place to form a zinc-bound water (Zn-H₂O) (Eq. 1). The second step of the reaction is the transfer of a proton from Zn-H₂O to the bulk solvent (B) to regenerate the Zn-OH⁻ for the next catalytic cycle, this is achieved via a network of ordered waters in the active site (proton wire) and a proton shuttling residue His64, at the entrance of the active site cavity [3–6]. This proton transfer (PT) occurs at the rate of 10⁶ s⁻¹ and is the rate-limiting step of the overall maximum velocity of catalysis [6, 7].

The high catalytic activity, ease of expression, and purification of HCA II has generated interest in its use in carbon sequestration systems to reduce the amount of atmospheric CO₂ emissions from the burning of fossil fuels [8, 9]. However, the denaturation of the enzyme at high temperatures and acidic pH is a resultant of the process, and therefore limits the usefulness of wild-type HCA II in these systems. As such, there is a wide interest in the bioengineering of a thermal stable HCA II variant that can withstand these harsh industrial settings [10, 11].

The rational design of thermal stable enzymes has proven to be a significant challenge in computational approaches because the proposed stabilizing elements such as core packing, surface electrostatics, and rigidity vary widely among different proteins [12–14]. Comparison of proteins isolated from thermophilic organisms to their mesophilic counterparts often reveals a sequence identity of 40 – 85% [15, 16], superimposable tertiary structure [17–23], and small differences (5 – 20 kcal mol⁻¹) in the free energy of stabilization (ΔG) between the folded states [24]. In addition, stability studies of enzyme variants revealed that differences in ΔG as small as 3.0 – 6.5 kcal mol⁻¹ can account for an increase in melting temperature (T_M) by 12 °C [25, 26]. These small energy differences are encouraging for the design of thermal stable variants via site-directed mutagenesis, but caution must also be taken as unforeseen effects on the catalytic activity, folding, and solubility of the enzyme can occur [27, 28].

One current working hypothesis is that hyperthermophilic enzymes are more rigid than their mesophilic homologues and have a lower entropy of unfolding [27]. This is supported by numerous experimental data including frequency domain fluorometry and anisotropy decay [29], hydrogen-deuterium exchange [30–32], and tryptophan phosphorescence [33]. Proline, which has limited conformation geometry within the main-chain of a polypeptide chain and restricts the conformation of the preceding residues [34], has the lowest entropy of all the amino acids. There are a number of examples of engineered thermophilic enzymes that have utilized the concept of proline incorporation with increased stability [34–37]. Residue positions Lys170 and Glu234 in HCA II are excellent candidates for proline incorporation because they are located in β-turns, and therefore naturally adopt the restricted φ angle of proline. Thus, these sites contain the required dihedral angles seen in the i+1 position for Type I and II β-turns [38].

Additionally, loop truncation of several mesophilic proteins have been shown to increase the T_M [39, 40]. HCA II has an extended loop region between residues 230–240 that is a good candidate for such a truncation. In the HCA II crystal structure this loop has a higher average C_α B-factor (thermal fluctuation) than the average for the enzyme and contains two solvent-exposed hydrophobic residues (Phe231 and Leu240) which have been previously shown to have detrimental effects on the thermal stability [11]. Chain annealing after deletion of the 230–240 loop would not be detrimental to the protein backbone, as the carbonyl of Leu229 would be sufficiently close to the amine of Met241 (2.8 Å) for ligation. Furthermore, the structural comparison of the hyperthermophilic CA isolated from the bacterium Sulfurihydrogenibium yellowstonense YO3AOP1 (SspCA) with HCA II revealed this loop to be absent in SspCA [41].

This study describes the rigidification of HCA II via proline substitutions and surface loop deletion. These elements have only minor effects on the thermal stability, but do have a significant impact on the catalytic activity of the enzyme.

Results

Differential Scanning Calorimetry

The thermograms for wild-type, K170P, E234P and Δ230–240 HCA II were obtained in triplicate at pH 7.8 before being averaged, buffer-subtracted and baseline corrected. The major transition peak for all variants displayed a single unfolding dynamic (Fig. 1). The transitions were calculated to be endothermic from Eq. 3 and were centered at the T_M, with a maximum heat capacity (C_P) occurring at the midpoint of the peak. A summary of the calculated T_M values is given in Table 1. The calorimetric enthalpy at the transition midpoint (ΔH^°_M) was calculated via integration of the area under the unfolding peak and normalized to the protein molar concentrations. The identity of the van’t Hoff enthalpies (ΔH^vH) to that of ΔH^°_M (~230 kcal mol⁻¹) confirmed that the unfolding process is a reversible two-state transition for all variants.

Figure 1.

Thermograms of the unfolding transition for A) wild-type; B) K170P; C) E234P; and D) Δ230–240 HCA II at pH 7.8. The buffer subtracted, baseline-corrected data are shown in black with the fit to the data shown in red.

Table 1.

Thermal Stability and Catalytic Measurements

Enzyme	TM (°C)a	kcatex/KeffS (M−1µs−1)b	kB (µs−1)c	pKa ZnOH−c	pKa His64c
wild-type HCA II	57.1 (± 0.1)	120 (± 5)	1.2 (±0.2)	6.9 (± 0.1)	7.2 (± 0.1)
K170P	55.7 (± 0.1)	63 (± 3)	1.4 (±0.3)	6.7 (± 0.1)	6.4 (± 0.1)
E234P	57.9 (± 0.1)	56 (± 2)	3.1 (± 0.6)	7.2 (± 0.1)	6.2 (± 0.1)
Δ230–240	58.9 (± 0.2)	66 (± 3)	1.0 (± 0.6)	6.7 (± 0.2)	5.8 (± 0.2)

^aDenaturation temperature as determined from Fig. 1 and Eq. 3.

^bCatalytic efficiency determined from the fit in Fig. 2A and Eq. 8

^cProton transfer rate and ionization constants determined from the fit in Fig. 2B and Eq. 9

There were only modest measured increases in the T_M for the E234P and Δ230–240 HCA II (T_M ≈ 58 and 59 °C, respectively; Figs. 1C and D) compared to wild-type HCA II (T_M ≈ 57 °C; Fig. 1A). Surprisingly, K170P HCA II, showed a slight lowering in stability with a T_M ≈ 56 °C (Fig. 1B). The minor increase of E234P and decrease of K170P HCA II T_Ms do not follow the value reported with a proline substitution at residue L240P HCA II, which increased the thermal stability to 60 °C [42].

Catalytic Activity

The pH profiles for the two rate constants, k_cat/K_M and R_H2O, were determined by measuring the exchange of ¹⁸O-label between CO₂ and water using mass spectrometry (Eqs. 6 and 7). The rates of catalyzed interconversion between CO₂ and HCO₃⁻, as measured by k_cat/K_M (Eq. 8) for K170P, E234P and Δ230–240 HCA II were all measured to be ~60 M⁻¹ µs⁻¹, an approximate 2-fold decrease compared to wild-type HCA II (Fig. 2A; Table 1). The measured rate constant for PT, k_B, from the ZnH₂O through the water network to His64 was determined by a fit from Fig. 2B curve to Eq. 9 and showed that K170P HCA II PT was comparable to that of wild-type HCA II (~1.4 µs⁻¹), whereas E234P had a ~2.5 fold increase in k_B (~3.1 µs⁻¹) (Table 1). Interestingly, the pH profile for Δ230–240 displayed a broad curve that is indicative of a double-ionization fit that describes a k_B rate that is similar to wild-type HCA II, but with larger uncertainty in the maximum rate (Table 1). All three variants revealed a pKa for the ZnOH⁻ similar to wild-type HCA II (~7.0), but the pK_a range for the proton shuttle residue, His64, is significantly different (5.8 – 6.4 for the variants compared to 7.2 for wild-type HCA II; Table 1).

Figure 2.

pH profiles for A) catalytic efficiency (k_cat/K_M) and B) proton transfer rate (R_H2O/[E]) for the HCA II variants. Wild-type is shown as a closed square, K170P as an open diamond, E234 as an open triangle and Δ230–240 HCA II as a closed circle.

X-ray Crystallography

The K170P and Δ230–240 HCA II crystallized in conditions similar to that of the wild-type HCA II [43], but the crystal growth took longer (1 – 3 months) and formed in the orthorhombic P2₁2₁2₁ space group, whereas wild-type HCA II crystals usually grow in one week and form in a monoclinic P2₁ space group. The K170P crystals grew using a protein concentration of 10 mg/mL (the same as wild-type), whereas Δ230–240 required a 5-fold higher concentration of 55 mg/mL. The change in space group of the K170P and Δ230–240 HCA II is most likely a direct consequence of a conformation change in the 230–240 loop in K170 HCA II and its deletion in the Δ230–240 HCA II, as in the wild-type HCA II monoclinic P2₁ structure this loop is involved in a crystallographic contact (Fig. 3A). The E234P variant crystallized in similar conditions, concentration, time, and space group as compared to wild-type HCA II, and also had no effect on the conformation of the 230–240 loop. The HCA II variants diffracted to a maximum resolutions of 1.35 Å for Δ230–240, 1.52 Å for E234P and 1.60 Å for K170P HCA II. A summary of the collected diffraction and final refinement statistics is shown in Table 2.

Figure 3.

Topology of the 230–240 loop in the HCA II variants. A) Cartoon view of wild-type HCA II (green) with the 230–240 loop highlighted in purple. The Zn²⁺ (magenta sphere) with the coordinating His residues and the proton shuttle residue, His64 shown in stick. B) Comparison of the 230–240 loop in the K170P (blue) to wild-type (green) HCA II. Side-chains are shown in stick and are as labeled. Residues G233 and G235 are not labeled for clarity. C) Comparison of the 230–240 loop in E234P (light purple) to wild-type (green) HCA II. Residues are as labeled with potential salt bridge interactions shown as a dashed line. D) Ligation between residues L229 and M241 in Δ230–240 (light pink) showing the deletion of the loop compared to wild-type HCA II (green) HCA II.

Table 2.

X-ray crystallographic data set and refinement statistics for K170P, E234P and Δ230–240 HCA II.

	K170P	E234P	Δ230–240
PDB Accession Number	4QK1	4QK2	4QK3
Wavelength (Å)	1.5418	1.5418	0.9177
Spacegroup	P212121	P21	P212121
Unit cell parameters (Å; °)	a = 42.0, b = 69.0, c = 73.6; β = 90	a = 42.2, b = 41.1, c = 71.9; β = 104.3	a = 42.1, b = 69.0, c = 74.2; β = 90
Resolution	20.0 – 1.60 (1.66 – 1.60)*	20.0 – 1.52 (1.57 – 1.52)	20.0 – 1.35 (1.40 – 1.35)
Total number of measured reflections	179331	259705	168087
Total number of unique reflections	27184	33910	44710
Rsyma(%)	10.4 (43.9)	8.2 (24.2)	12.0 (48.0)
I / σ I	11.4 (3.2)	25.8 (4.4)	11.2 (2.0)
Completeness (%)	93.9 (99.8)	93.4 (89.1)	91.9 (93.8)
Redundancy	6.6 (6.8)	7.7 (7.7)	3.8 (3.3)
Rcryst (%)	17.7	15.9	21.7
Rfree (%)	22.3	17.8	25.0
Residue Number	256	257	246
Number of water molecules	195	275	167
r.m.s.d.: Bond lengths (Å); angles (°)	0.011; 1.486	0.008; 1.215	0.010; 1.440
Coordinate error (Å)	0.3	0.2	0.2
Ramachandran statistics (%): Most favored; allowed	94.9; 5.1	96.9; 3.1	94.6; 5.4
Average B-factors (Å2): All; main-; side-chain; solvent	28.4; 25.4; 30.2; 32.9	17.0; 13.0; 17.4; 29.5	21.9; 17.6; 23.6; 34.5

^aR_sym is defined as ∑_hkl ∑_i |(I_i(hkl)−<I_i(hkl) >|/ ∑_hkl∑_iI_i(hkl) × 100, where I_i(hkl)is the intensity of an individual reflection and <I_i(hkl) > is the average intensity for this reflection; the summation is over all intensities.

^bR_cryst = (∑|F_o − F_c|/∑ |F_o|) × 100

^cR_free is calculated in the same way as R_cryst except it is for data omitted from refinement (5% of reflections for all data sets).

^*Values in parentheses are for the highest resolution shell.

The initial mF_o – DF_c omit maps revealed positive density (>4σ) around the K170 and E234 amino acid sites and after building as proline side chains, the refined 2mF_o – DF_c maps unambiguously confirmed the incorporation of a proline residues at positions 170 and 234, with a maximum electron density of 0.32 and 0.46 e/ų, respectively. Likewise, the deletion of the surface loop between residues 230 and 240 was seen as negative density (>4σ) in the initial mF_o – DF_c omit map in the region between Leu229 and Met241, confirming the Δ230–240 HCA II variant. After the deletion of the loop in the model and subsequent rounds of refinements, excellent electron density in the final 2mF_o – DF_C map for the ligation of Leu229 and Met241 was observed with a maximum electron density of 0.55 e/ų, comparable to that of the surrounding peptide backbone.

Superimposition of the wild-type HCA II (PDB: 3KS3; [43]) onto E234P HCA II (Fig. 3C) revealed an r.m.s.d. of main-chain of 0.2 Å, indicating no change in the main-chain conformation. However, K170P HCA II (Fig. 3B) exhibited an alternative 230–240 loop main-chain conformer with a mean r.m.s.d. of 2.2 Å. This conformational loop change is most likely a direct consequence of the K170P substitution, as the basic lysine residue is involved in electrostatic interactions with the 230–240 acidic loop in wild-type HCA II. This would also explain why the observed space group changes from monoclinic to orthorhombic. All three structures, however, displayed an overall topology similar to that of the wild-type HCA II with an r.m.s.d. of ~0.2, 0.3 and 0.7 Å for the E234P, Δ230–240 and K170P HCA II C_α atoms, respectively. The incorporation of a proline residue at position 234 increased the overall rigidity of the 230–240 loop by ~9% compared to the wild-type HCA II structure as measured by the relative B-factors and this correlates well to the enhanced thermal stability (Fig. 1; Table 1). For comparison, the relative B-factors of the C_α backbone in the 230–240 loop for the K170P HCA II variant increased by ~33% compared to wild-type HCA II which also correlates to its apparent increased flexibility and loss in thermal stability. The proline substitution at residue 170 did have a slight effect on the rigidity (~2% decrease in the relative C_α backbone B-factors compared to wild-type) in the region between residues 167 and 173.

Comparison of the active site residues and water molecules involved in the PT network for K170P and E234P HCA II revealed very little deviation in their positioning relative to wild-type HCA II, and therefore provided no structural evidence for their altered catalytic efficiencies and PT rates. The Δ230–240 HCA II active site water molecules W1 and W2 were in approximate identical locations (0.2 Å) compared to wild-type (Fig. 4). There was a significant shift in the positioning of W3B (2.8 Å), which interacts with Asn62 and Asn67 in wild-type HCA II, to form interactions with Gln92 and W4 in the Δ230–240 HCA II active site. W4 was also repositioned by 2.2 Å resulting in loss of a hydrogen bond with Gln92 as compared to wild-type HCA II. Interestingly, the shifts seen for W1 and W2 in the Δ230–240 HCA II active site places W2 too far away from His64 for a potential hydrogen bond (3.9 Å; Fig. 5).

Figure 4.

Active site water network configuration in Δ230–240 (red spheres) overlaid with the wild-type (light pink spheres) HCA II. Potential hydrogen bond interactions for Δ230–240 are shown as black dashed lines whereas the white dashed lines represent the hydrogen bonds observed in wild-type HCA II Residues and water molecules are as labeled.

Figure 5.

Schematic of the proton transfer network for A) wild-type and B) Δ230–240 HCA II. Hydrogen bond interactions (not drawn to scale) are depicted as arrows with the distance between atoms label in Å.

Discussion

Previous studies have proposed that thermophilic enzymes owe their stability in part to the overall compactness and rigidity compared to their mesophilic homologues. For HCA II, this was shown to be true when a proline at position 240, L240P, was engineered into the enzyme and enhanced the thermal stability (ΔT_M ≈ 3 °C) compared to wild-type HCA II [11, 42]. Additionally, the recently determined X-ray crystallographic structure of SspCA [41] implied that the absence of the 230–240 loop (HCA II numbering) may be linked to its extreme thermal stability (T_M > 100 °C). The modest ~2 °C increase in melting temperature of Δ230–240 HCA II does support the hypothesis that thermophilic proteins display a more compact topology. However, this cannot account for the extreme thermal stability reported for SspCA (T_M > 100 °C; [41]), but it may arise from other elements including the incorporation of ionic clusters on the surface and an extensive dimeric interface (HCA II being a monomer). Furthermore, the increase of the Δ230–240 HCA II stability is unremarkable when compared to previously reported HCA II variants which increased the T_M by decreasing the surface hydrophobicity (ΔT_M ≈ 7 °C) [11], and the incorporation of a conserved disulfide bridge (ΔT_M ≈ 14 °C) [10].

The DSC studies of the E234P variant revealed only a modest 1 – 2 °C increase of T_M compared to wild-type HCA II (Fig. 1; Table 1). The enhanced thermal stability is consistent with previous studies [11, 42]), but may imply that the greatest stabilizing effect may arise if the target residue is nonpolar. In contrast, the K170P variant has a T_M compared to wild-type HCA II (Fig. 1; Table 1) despite a decreased thermal fluctuation in the 167–173 loop, as determined from B-factor analysis. The loss in thermal stability of K170P HCA II can be associated with the increase in flexibility of the 230–240 loop via loss of the tethering to the acidic loop by the basic residue (Fig. 3B). Site-directed mutagenesis of Lys172 may verify this hypothesis as it seems to provide potential electrostatic interactions with the 230–240 loop in addition to Lys170. Taken together, while it seems that the thermal stability of HCA II can be enhanced via rigidification and compression of the accessible surface area, there also seems to be some dependence on the surface polarity.

Measurement of the catalytic activity via ¹⁸O mass spectrometry of the HCA II variants revealed a two-fold decrease in k_cat/K_M compared to wild-type HCA II (~60 compared to 120 M⁻¹ µs⁻¹, respectively) with varying rates in PT (Fig. 2; Table 1). Additionally, the pK_a of the active site was measured and shown to be altered in the variants as compared to wild-type HCA II, presumably as a direct effect of either loss in the 230–240 loop acidity in the E234P and Δ230–240 variants or reconfiguration of the loop around the peripheral of the active site with the K170P variant. These altered ionization constants for His64 could explain the increased PT rates for E234P HCA II, as a deprotonated His64 would be better suited to accept a proton from the water network. The lowered catalytic efficiency of the HCA II variants may be associated with these lowered pK_a values for His64 by having an effect on the binding affinity and/or coordination of substrate molecules in the active site. While this effect is minimum in the positioning of the water molecules within the active site of the K170P and E234P variants, deletion of the acidic loop between residues 230 and 240 resulted in a distorted PT water network that included shifts in the positioning of W1, W2, W3B and W4, as well as the inclusion of an additional water molecule (W2A; Fig. 4). The W2A molecule shortens the hydrogen bond distance between W2 and W3A from 3.4 Å in wild-type HCA II (Fig. 5A) to ~2.8 Å in Δ230–240 HCA II (Fig. 5B). PT between W3A and His64 can then potentially occur at a distance (3.2 Å) similar to that observed in wild-type HCA II (Fig. 5). As seen in the R_H2O pH profile (Fig. 2B), the rate of PT in the Δ230–240 HCA II variant has less of a dependence on the pH of the solvent and may reconfigure its PT water network in accordance with its environment. Further crystallographic studies at varying pH are needed to elucidate this unique water network.

Conclusions

The bioengineering of enzymes to meet specific criteria can be a valuable asset in a variety of industrial and medical systems. The ever increasing demand for a carbon sequestration agent to help combat global warming and the need for a reliable biomedical catalyst has fueled research into the rational design of a thermal stable variant of HCA II without detriment to high catalytic activity. Through superimposition of thermophilic proteins onto their mesophilic counterparts, several elements have been proposed to provide stability including a more rigid and compact surface, increased ionic networks on the enzyme surface and the inclusion of disulfide bridges and aromatic clusters within the enzyme core [12–14, 44, 45]. The extent to which these stabilizing elements contribute to the overall stability, however, fluctuates among different proteins. This study suggests that the stability of HCA II can be enhanced by rigidification of surface loops via incorporation of proline residues as well as increasing the surface compactness via loop deletion. The positioning of these alterations, however, may have unforeseen effects on the catalytic activity of HCA II as the acidic loop between residues 230 and 240 seems to modulate the pK_a of the active site. Reduction of the electrostatic potential in this loop via the E234P substitution and reconfiguration of the loop through the K170P substitution resulted in decreased catalytic efficiency and altered PT rates whereas deletion of the 230–240 loop resulted in a complete reorganization of the active site PT water network. The successful biophysical analysis of HCA II variants containing the aforementioned thermal stabilizing elements could provide valuable insight into protein stability and promote significant developments in bioengineering computational techniques.

Materials & Methods

Protein Expression and Purification

HCA II cDNA containing the K170P and E234P mutations were prepared from an expression vector containing the enzyme coding region [46] via site-directed mutagenesis using the Stratagene QuikChange II kit and primers designed from Invitrogen. The plasmid cDNA expressing the loop deletion between residues 230 and 240 in HCA II (Δ230–240 HCA II) was created utilizing the PCR-mediated DNA deletion method [47]. In short, two primers (Primers A and B) were designed such that Primer A is the reverse complement of a sequence corresponding to 20 bases upstream of the sequence to be deleted followed by a reverse complimentary sequence 20 bases downstream of the deletion site. Primer B was designed in the same fashion but corresponding to the complementary strand; that is, the sequence of Primer B was identical to the plasmid primary lacking the sought deletion. The corresponding cDNA for each variant was transformed into E. coli XL1-Blue super-competent cells, which were then confirmed by DNA sequencing of the entire coding region.

The verified cDNA for each HCA II variant was transformed into E. coli BL21(DE3) cells in 1L of 2 × Luria broth containing ~0.1 mg/mL ampicillin and grown at 37 °C until a turbidity of ~0.6 measured at 600 nm was reached. Protein production was induced via the addition of ~0.1 mg/mL isopropyl β-D-1-thiogalactopyranoside (IPTG) and metal inclusion in the enzyme active site was insured via addition of ~1 mM zinc sulfate (final concentrations). The cells were incubated for an additional three hours and harvested by centrifugation for 12 minutes at 4000 rpm in a JA-10 rotor. The resulting pellet was suspended in 200 mM sodium sulfate, 100 mM Tris-HCl, pH 9.0 and lysed by addition of hen egg white lysozyme. Cellular debris was removed via addition of DNaseI followed by centrifugation for 75 mins at 15000 rpm in a JA-20 rotor. The resulting lysate was filtered using a 0.8 µm syringe filter (Amicon) and purified on an affinity column containing an agarose resin coupled with p-(aminomethyl)-benzene-sulfonamide, a tight-binding inhibitor of HCA II [48]. The bound HCA II variants were then eluted with 400 mM sodium azide, 100 mM Tris-HCl, pH 7.0 followed by concentration and buffer-exchanging in 50 mM Tris-HCl, pH 7.8 using Amicon Ultra centrifuge filters (10 kDa MWCO) at 8000 rpm in 12 min integrals using a JA-20 rotor.

Differential Scanning Calorimetry

The DSC experiments were performed to assess the thermal stability of the HCA II variants under near-physiological conditions using a VP-DSC calorimeter (Microcal, Inc., North Hampton, MA) with a cell volume of ~0.5 mL. The HCA II samples (6–10 µM) were extensively buffer-exchanged into 50 mM Tris-HCl, pH 7.8 and then degassed, while stirring, at 16 °C for 20 minutes prior to data collection. The DSC thermograms were collected in triplicate from 20 to 100 °C with a scan rate of 60 °C/h. The calorimetric enthalpies of unfolding were calculated by integrating the area under the peaks in the thermograms after adjusting the pre- and post-translation baselines. The thermograms were fit to a two-state reversible unfolding model to obtain van't Hoff enthalpies (ΔH^vH) of unfolding.

The HCA II variant melting temperature (T_M) values were obtained from the midpoints of the DSC curves, indicating a two-state transition. The difference in Gibbs free energy (ΔG°) at a given temperature, T, was calculated via [49]:

$$\mathrm{\Delta }\mathrm{G}°(\mathrm{T})=\mathrm{\Delta }{\mathrm{H}°}_{\mathrm{M}}(1-\mathrm{T}/{\mathrm{T}}_{\mathrm{M}})+\mathrm{\Delta }{\mathrm{C}}_{\mathrm{P}}((\mathrm{T}-{\mathrm{T}}_{\mathrm{M}})-\mathrm{T}\text{ln}(\mathrm{T}/{\mathrm{T}}_{\mathrm{M}}))$$ (3)

Where ΔH°_M is the calorimetric enthalpy at T_M and ΔC_P is the observed change in heat capacity between the folded and unfolded states. The denaturation enthalpies (ΔH°) and entropies (ΔS°) were calculated at a given temperature using the Kirchoff's law equations [43]:

$$\mathrm{\Delta }\mathrm{H}°(\mathrm{T})=\mathrm{\Delta }{\mathrm{H}°}_{\mathrm{M}}+\mathrm{\Delta }{\mathrm{C}}_{\mathrm{P}}(\mathrm{T}-{\mathrm{T}}_{\mathrm{M}})$$ (4)

$$\mathrm{\Delta }\mathrm{S}°(\mathrm{T})=\mathrm{\Delta }{\mathrm{S}°}_{\mathrm{M}}+\mathrm{\Delta }{\mathrm{C}}_{\mathrm{P}}\text{ln}(\mathrm{T}-{\mathrm{T}}_{\mathrm{M}})$$ (5)

Thermograms for all samples were obtained in triplicate and then averaged to obtain the final profile, which was then used for reference subtraction and data analysis.

Kinetic Studies

The ¹⁸O exchange method is based on mass spectrometric measurements using a membrane inlet of the depletion ¹⁸O from CO₂ [50]. The isotopic content of CO₂ in solution is measured when it passes across a membrane and into an Extrel EXM-200 mass spectrometer. The measured variable is the atom fraction of ¹⁸O in CO₂. The first step of catalysis has a probability of reversibly labeling the ZnOH⁻ with ¹⁸O (Eq. 6). During the next step the ¹⁸OH⁻ can be protonated and results in the release of H₂¹⁸O to the bulk solvent where it is essentially infinitely diluted by H₂¹⁶O (Eq. 7). In this process, His64 acts as a proton shuttle [5].

$$\mathrm{H}\mathrm{C}\mathrm{O}{\mathrm{O}}^{18}{\mathrm{O}}^{-}+\mathrm{E}\mathrm{Z}\mathrm{n}{\mathrm{H}}_2\mathrm{O}\rightleftarrows \mathrm{E}\mathrm{Z}\mathrm{n}\mathrm{H}\mathrm{C}\mathrm{O}{\mathrm{O}}^{18}{\mathrm{O}}^{-}\rightleftarrows \mathrm{C}\mathrm{O}\mathrm{O}+\mathrm{E}\mathrm{Z}{\mathrm{n}}^{18}\mathrm{O}{\mathrm{H}}^{-}$$ (6)

$${\mathrm{H}}^{+}\mathrm{H}\mathrm{i}\mathrm{s}64-\mathrm{E}\mathrm{Z}{\mathrm{n}}^{18}\mathrm{O}{\mathrm{H}}^{-}+{\mathrm{H}}_2\mathrm{O}\rightleftarrows \mathrm{H}\mathrm{i}\mathrm{s}64-\mathrm{E}\mathrm{Z}\mathrm{n}{{\mathrm{H}}_2}^{18}\mathrm{O}\rightleftarrows \mathrm{H}\mathrm{i}\mathrm{s}64-\mathrm{E}\mathrm{Z}\mathrm{n}{\mathrm{H}}_2\mathrm{O}+{{\mathrm{H}}_2}^{18}\mathrm{O}$$ (7)

The ¹⁸O-exchange method obtains two different rates at chemical equilibrium [50]: R₁, which is the rate of exchange of CO₂ and HCO₃⁻ (Eq. 8); and R_H2O, which is the rate of release of H₂¹⁸O from the enzyme (Eq. 9). In Eq. 8, k_cat^ex is the rate constant for maximal conversion between substrate and product while K_eff^S is the effective binding constant of substrate ([S] is the concentration); [S] can be either CO₂ or HCO₃⁻ depending on the direction of the reaction. The ratio expressed in Eq. 8 of k_cat^ex/K_eff^S is in principle the same as k_cat/K_M obtained under steady state conditions.

$${\mathrm{R}}_1/[\mathrm{E}]={{\mathrm{k}}_{\text{cat}}}^{\text{ex}}[\mathrm{S}]/({{\mathrm{K}}_{\text{eff}}}^{\mathrm{S}}+[\mathrm{S}])$$ (8)

In the second step of catalysis, the rate R_H2O is the part of ¹⁸O exchange that is dependent on the rate of proton transfer from His64 to the labeled ZnOH⁻ (i.e. in the dehydration direction) [5]. Eq. 9 shows the relationship between k_B, the rate constant for proton transfer to ZnOH⁻ and (K_a)_donor and (K_a)_ZnH2O that are the ionization constants for the proton donor and ZnH₂O, respectively.

$${\mathrm{R}}_{\mathrm{H}2\mathrm{O}}={\mathrm{k}}_{\mathrm{B}}/[[1+{({\mathrm{K}}_{\mathrm{a}})}_{\text{donor}}/[{\mathrm{H}}^{+}]][1+[{\mathrm{H}}^{+}]/{({\mathrm{K}}_{\mathrm{a}})/}_{\mathrm{Z}\mathrm{n}\mathrm{H}2\mathrm{O}}]]$$ (9)

All enzyme kinetic measurements were done at 25 °C in the absence of buffer using a total substrate concentration (all species of CO₂) of 25 mM. Kinetic constants and ionization constants shown in Eqs. 8 and 9 were determined through nonlinear least squares methods (Enzfitter, Biosoft).

Crystallization and Diffraction Collection

Crystal trays were prepared for the K170P and E234P HCA II variant with a 1:1 ratio of the reservoir solution and the protein sample (10 mg/mL) in 5 µL drops utilizing the hanging drop vapour diffusion method. The reservoir solutions contained 500 µL of 1.4 M sodium citrate, 50 mM Na₂HPO₄, pH 8.0 for the K170P HCA II variant and 1.4 M sodium citrate, 50 mM Tris-HCl, pH 7.8 for the E234P variant. Crystal formation for K170P HCA II appeared after 3 months, whereas crystals for E234P HCA II appeared within one week. Crystal trays were prepared for Δ230–240 HCA II loop deletion variant in 1:1 ratio of the reservoir solution containing 1.6 M sodium citrate, 100 mM Tris-HCl, pH 7.8 and protein sample at 50 mg/mL in 5 µL drops utilizing the hanging drop vapour diffusion method. Crystal formation occurred within one week.

Diffraction data for K170P and E234P HCA II were collected on an in-house Rigaku R-Axis IV⁺⁺ image plate detector with a RU-H3R rotating Cu anode (K_α = 1.5418 Å) operating at 50 kV and 22 mA. The X-rays were focused using Osmic optics, followed by a helium-purged beam path. The crystal-to-detector distance was 80 mm for K170P and E234P HCA II. Each image was collected for five minutes with 1° oscillations. HKL2000 [51] was used to integrate, merge and scale the data sets. K170P HCA II crystallized in the orthorhombic space group P2₁2₁2₁ whereas E234P HCA II crystallized in the monoclinic space group P2₁. The crystals diffracted to 1.60 and 1.52 Å resolution for K170P and E234P, respectively. The diffraction and refinement statistics for K170P and E234P HCA II are summarized in Table 2.

Diffraction data for Δ230–240 HCA II was collected on the F1 beamline at Cornell High Energy Synchrotron Source (CHESS F1; λ = 0.9177 Å) on an ADSC Q-270 detector using the microfocused beam. The crystal-to-detector distance was 150 mm. Each image was collected for 10 seconds with 0.5° oscillations. HKL2000 [51] was used to integrate, merge and scale the data sets. Δ230–240 HCA II crystallized in the orthorhombic space group P2₁2₁2₁ and diffracted to a h 1.35 Å resolution. The diffraction and refinement statistics for Δ230–240 HCA II are summarized in Table 2.

Phase Determination and Structure Refinement

Initial phases for K170P and E234P HCA II were calculated using molecular replacement via Phaser [52] using the wild-type HCA II structure, with the residues at 170 and 234 replaced with alanine, respectively (PDB: 3KS3; [43]). 5% of the reflections were set for R_free calculations. Initial phases for Δ230–240 HCA II variant were determined using wild-type HCA II. PHENIX.REFINE [53] was used in cycles of restrained refinement of the molecular model, alternating with manual building using COOT [54]. Initial mF_o - DF_c phase maps revealed positive density at position 170 and 234 sidechain deletions, while the Δ230–240 loop deletion variant revealed negative density for the deleted residues 229 and 241. Subsequent refinements showed excellent electron density in the 2mF_o - DF_c maps for the K170P and E234P HCA II (Fig. 1A and B) as well as for the newly formed peptide linkage in Δ230–240 HCA II. The final R_cryst and R_free values are shown in Table 2. MOLPROBITY [55] was used to assess the quality of the final model. All structural figures were created in PyMOL [56].

B-factor Analysis

The thermal fluctuations of the loop regions encompassing residues 167–173 and 230–240 in all variants were compared via normalization to the corresponding residue with the highest C_α B-factor in each region. The B-factors of the remaining residues were converted to the percent of rigidification compared to the most flexible residue and the mean percentage was then calculated for each variant. These rigidification percentages of each variant was compared to HCA II in either the monoclinic spacegroup (PDB: 2ILI; [57]) or the orthorhombic spacegroup (PDB: 1UGG; [58]) for E234P and K170P HCA II, respectively, to minimize B-factors differences arising from resolution and spacegroup dissimilarities.

Abbreviations

CO₂

carbon dioxide

C_p

specific heat

DSC

differential scanning calorimetry

H⁺

proton

HCA II

human carbonic anhydrase II

HCO₃⁻

bicarbonate

SspCA

Sulfurihydrogenibium yellowstonense YO3AOP1 carbonic anhydrase

T_M

melting temperature

Zn-H₂O

zinc-bound water

Zn-OH⁻

zinc-bound hydroxide

ΔG

Gibb’s free energy