The first step was to determine how Cd²⁺ interacts with the C1B-C2 domain using UV-vis absorption spectroscopy. It is well established that thiolate-Cd²⁺ charge transfer bands have characteristic wavelengths at around ∼240 nm (Busenlehner et al., 2001; Habjanič et al., 2020). The C1B domain has six cysteine residues, all of which are involved in coordinating the structural Zn²⁺ ions (Figure 1A). C2 is cysteine-free, but can bind Cd²⁺ with high affinity through the vacant oxygen-rich sites formed by the aspartate carboxyl groups and the carbonyl oxygens of W247 and M186 (Morales et al., 2013a). Thus, the presence of thiolate-Cd²⁺ charge transfer bands upon C1B-C2 treatment with Cd²⁺ can only originate from Cd²⁺ coordinating Cys residues of C1B.

Addition of increasing amounts of Cd²⁺ to C1B-C2 resulted in significant spectral changes (Figure 1B). Based on the C2-only control experiment with Cd²⁺ (inset of Figure 1B), these changes can only be attributed to the C1B-Cd²⁺ interactions. The difference UV-Vis spectra, where the protein contribution to the absorbance is subtracted out, clearly shows the build-up of a shoulder near λ = 270 nm (Figure 1C). The wavelength range is consistent with the position of thiolate-Cd²⁺ charge transfer bands observed in other studies (Busenlehner et al., 2001; Habjanič et al., 2020). Based on this information and previous work on the Zn²⁺-containing proteins with Cys-rich sites (Wang et al., 2005; Chakraborty et al., 2011; Malgieri et al., 2011), we conclude that Cd²⁺ forms coordination bonds with the cysteine residues of C1B, even in the presence of Cd²⁺-sequestering C2.

Two scenarios are possible: Cd²⁺ can either eject and substitute for Zn²⁺, or Cd²⁺ can peripherally coordinate cysteines without displacing Zn²⁺, forming a binuclear metal cluster similar to that observed in the GAL4 transcription factor (Pan and Coleman, 1990). To distinguish between these two scenarios, we used a highly selective Zn²⁺ fluorophore, FluoZin-3. Four molar equivalents of Cd²⁺ were added to the C1B-C2 domain in the presence of FluoZin-3, and the time-dependent fluorescence intensity was monitored at 516 nm. We observed a steady increase in the fluorescence intensity, indicating that Zn²⁺ is being displaced from the protein as a result of Cd²⁺ treatment (Figure 1D, red trace). There was no time-dependent increase in fluorescence for an identical experiment conducted in the absence of externally added Cd²⁺ (Figure 1D, blue trace), indicating that Fluozin-3 alone cannot strip Zn²⁺ off C1B. Collectively, these experiments show that Cd²⁺ successfully ejects Zn²⁺ from C1B and forms coordination bonds with cysteines.

While the UV-vis data show that Cd²⁺ is displacing Zn²⁺ from C1B they do not contain any site-specific information. We used solution NMR spectroscopy to gain insight into how Cd²⁺ interacts with sites 1 and 2 of C1B (see Figure 1 for site definitions). The site-specific information was obtained by collecting 2D [¹⁵N, ¹H] HSQC spectra of [U-¹⁵N] enriched C1B^Zn in the absence and presence of Cd²⁺. Each N-H group in C1B^Zn gives rise to a cross-peak in the 2D NMR spectra that we assigned in our previous work (Figure 2A, red spectrum) (Stewart et al., 2011). Upon addition of Cd²⁺, we observed an appearance of a new subset of well-dispersed C1B cross-peaks (Figure 2A, black spectrum). We were able to assign this subset to specific Cd/Zn C1B states based on their relative peak intensities and the chemical shifts of the refolded C1B^Cd (vide infra). From the spectral overlay, it is evident that the N-H resonances of many C1B residues, particularly those coordinating Zn1 and Zn2, experience large chemical shift perturbations upon C1B binding Cd²⁺.

In addition to native C1B^Zn, there are three other possible Cd/Zn protein states: C1B^Cd, C1B^Zn/Cd, and C1B^Cd/Zn that can co-exist in solution. The N-H groups of three residues in C1B: His140, Ile145, and Val147 show four cross-peaks each (Figure 2B) and serve as direct evidence for the formation of the all-Cd and Zn/Cd mixed C1B species. Moreover, the distinct chemical shifts of the four cross-peaks enable the calculation of the relative affinities of Cd²⁺ to each metal ion coordination site, using the procedures described in the Materials and Methods section. The relative affinity data presented in Table 1 show that: (i) Cd²⁺ has a ∼2-fold and ∼1.6-fold higher affinities than Zn²⁺ for the C1B sites 1 and 2, respectively; and (ii) relative Cd²⁺ affinity for either site does not depend on the chemical identity of the metal ion, Cd²⁺ or Zn²⁺, that occupies the other site (i.e. for a given site the χ and μ values are essentially identical). We conclude that both thiol-rich coordination sites in C1B are reactive with respect to Cd²⁺ substituting for the native Zn²⁺ ion.

In the PKCα regulatory region, C1B is adjacent to the C2 domain. C2 is metal-ion free in the inactive state of the kinase, but binds Ca²⁺ that is released as a result of the signaling events preceding PKCα activation. The Ca²⁺ binding site is formed by the Ca²⁺- and membrane-binding loops or CMBLs (Figure 1A). To determine the effect of the C2 domain on the C1B-Cd²⁺ interactions, we compared the [¹⁵N,¹H] HSQC spectra of C1B-C2 in the absence and presence of 2 molar equivalents of Cd²⁺. We observed the same signatures of Zn²⁺ replacement as in the isolated C1B domain, including the presence of four cross peaks for Ile145 and Val147 (Figure 3A). Overall, there is an excellent correlation between the chemical shift perturbations due to Cd²⁺ binding for isolated C1B and C1B in the context of its neighboring C2 (Figure 3B, inset). The full chemical shift perturbation (CSP) plot shows that not only C1B resonances are affected by interactions with Cd²⁺, but also the CMBLs of C2 (Figure 3B). We previously demonstrated that the isolated C2 domain can bind Cd²⁺ with high affinity (K_d < 1 μM) through the loop regions (Morales et al., 2013a). Collectively, these data indicate that Cd²⁺ binds simultaneously to both C1B and C2 domains and that the thiol-rich C1B Cys₃His sites can effectively compete for Cd²⁺ with the C2 oxygen-rich sites.

It is evident from the chemical shift dispersion in the 2D spectra that C1B remains folded upon incorporating Cd²⁺ (Figure 2 and Figure 3A). To test if C1B^Cd retains its function, we conducted NMR-detected binding experiments between C1B^Cd and a tumor-promoting agent, phorbol-12,13-dibutyrate (PDBu, Figure 4A). PDBu is an extremely potent exogenous agonist of PKC that binds specifically to C1 domains and drives their membrane insertion as part of the PKC activation sequence. These properties have made PDBu the most commonly used agonist (Katti and Igumenova, 2021) in the PKC field to assess the C1 domain functional competency. To generate C1B^Cd as the dominant species in solution, C1B^Zn was denatured and refolded in the presence of Cd²⁺. The 2D [¹⁵N,¹H] HSQC spectrum of the refolded C1B^Cd showed distinct chemical shifts compared to those of C1B^Zn (Figure 4B), but superimposed exactly onto the spectrum of the Cd²⁺-bound species that were formed as a result of C1B^Zn treatment with Cd²⁺ (Figure 2A).

PDBu is an extremely hydrophobic ligand that requires a membrane-mimicking environment to form a soluble complex with C1 domains. To provide such an environment, we used the DPC/DPS mixed micelle system that supports the C1 ligand-binding function (Stewart et al., 2011; Stewart et al., 2014) and faithfully reproduces the outcomes of in-cell experiments. Upon addition of PDBu and mixed micelles to C1B^Cd, we observed dramatic changes in the NMR spectrum (Figure 4B). Several residues, such as Ser111, Gly124, Leu125, and Ile126 experienced significant chemical shift perturbations upon the formation of the ternary C1B^Cd-PDBu-micelle complex. The CSP plot comparing the complex with the apo state showed that the changes are localized to the C1B membrane-binding loop regions, which is responsible for capturing the ligand in the membrane environment (Figure 4C). This CSP pattern is essentially identical to that observed for the native C1B^Zn protein upon PDBu binding in micelles (Stewart et al., 2011). Because NMR chemical shifts are exquisitely sensitive to the electronic environment of the reporting nuclei, we conclude that C1B^Cd interacts with PDBu and partitions into micelles in a manner identical to that of the native C1B^Zn.

To investigate the site-specific kinetics of Zn²⁺ replacement with Cd²⁺, we used SOFAST HMQC experiments to monitor the build-up of the Cd²⁺-bound C1B species. The population in % was calculated as the ratio of the N-H cross-peak intensities of the Cd²⁺-bound C1B, I_Cd, and the combined peak intensities I₀ = I_Cd + I_Zn. The data were plotted as the mean of the I_Cd/I₀ values for a subset of residues (listed in the Methods section) that report on Cd²⁺ binding to either site 1 or site 2. The kinetics data shown in Figure 5A revealed that sites 1 and 2 differ with respect to their kinetic behavior.

Site 2 is more reactive towards Cd²⁺, reaching the Cd²⁺-bound population of 53% within the first 15 min of the experiment. This exceeds the equilibrium value by ∼10%, and the site 1 population by 17%. As shown on the 3D structure of the C1B domain in Figure 5B, Zn²⁺ at site 2 brings the termini of C1B together by coordinating His102 at the N-terminus and Cys151 at the C-terminus. This part of the protein has a relatively high degree of solvent exposure and is therefore readily accessible to Cd²⁺. Another distinct feature of site 2 is the presence of a reactive Cys residue, Cys151, which serves as the entry point for the reactive oxygen species that activate PKCα in a process involving Zn²⁺ release. The structural dynamics of site 2, associated with the loss of Cys151 coordination bond with Zn²⁺ (Stewart and Igumenova, 2012), is likely to be another factor that makes site 2 susceptible to Cd²⁺ interactions. Under the conditions of our experiment, the system reached equilibrium within 1 hour. At equilibrium, the Cd²⁺ population of site 1 is higher than that of site 2, fully consistent with the pattern of relative Cd²⁺ affinities (Table 1). Together, the data of Figure 1D, 5(A), and Table 1 show that Cd²⁺ binding accompanied by Zn²⁺ ejection is a slow process, and that sites 1 and 2 are non-equivalent kinetically and thermodynamically.

The Cd²⁺ stock solutions were prepared by dissolving Cd(NO₃)₂·4H₂O (>99% purity, Sigma-Aldrich) in the appropriate buffer. Unless indicated otherwise, the experiments were conducted in the “MES buffer” comprising 10 mM 2-(N-morpholino)ethanesulfonic acid (MES) at pH 6.0 in HPLC-grade water (Avantor), 150 mM KCl, and 1 mM tris(2-carboxyethyl) phosphine (TCEP). The buffers were passed through the Chelex^® 100 (Sigma-Aldrich) column to remove residual divalent metal ions.

The DNA sequences encoding C1B-C2 (residues 100–293), isolated C1B (residues 100–152) or C2 (residues 155–293) of PKCα (M. musculus for C1B-C2 and C1B; R. Norvegicus for C2) were amplified by PCR using the cDNA clone of PKCα (Open Biosystems) as a template and cloned into the pET-SUMO vector (Invitrogen). Isolated C1B, C2, and C1B-C2 were expressed and purified as described previously (Morales et al., 2011; Stewart et al., 2011; Cole et al., 2019). [U-¹⁵N, 75%-²H]-enriched C1B-C2 and [U-¹⁵N]- or [U-¹³C, ¹⁵N]-enriched C1B were used for the NMR experiments.

UV-vis spectra were collected on a Beckman DU 640 spectrophotometer. 25 μM protein (C1B-C2, C2, or C1B) solution or MES buffer (for metal ion-only reference experiments) were placed in the sample cuvette; the reference cuvette always contained metal ion-free MES buffer. Cd²⁺ was added stepwise from the corresponding stock solutions to the sample cuvette. The samples were incubated for 1 h prior to the start of the measurements. The post-acquisition processing included the subtraction of the free Cd²⁺ spectra from the spectra of protein-containing samples. To eliminate contribution of protein-only absorption bands, the difference spectra were generated by subtracting the spectrum of the apo protein from the spectra of the metal-ion-containing protein. All spectra were corrected for dilution prior to subtraction.

[U-¹⁵N]-enriched C1B was dissolved in 6 M guanidine hydrochloride (Acros Organics) and the “refolding buffer” comprising 20 mM MES at pH 6.0 and 1 mM TCEP. The final protein concentration was between 15 and 35 μM during the denaturation step. The refolding was conducted in three dialysis steps, all of them carried out in the refolding buffer: (1) against 8 M urea at room temperature, for 8 h; (2) against 1.5 M urea and 100 μM Cd(II) nitrate at 4°C, overnight; and (3) against urea-free buffer at 4°C, for 3 days to ensure complete removal of urea. The refolded protein was concentrated in a Vivaspin^® spin concentrator with a 3 kDa cut-off and subsequently exchanged into an “NMR buffer” (10 mM MES at pH 6.0, 150 mM KCl, 1 mM TCEP, 0.02% NaN₃, and 8% (v/v) D₂O using a Midi-Trap G25 desalting column (GE Healthcare).

All proteins were concentrated and buffer exchanged using 10 kDa (C1B-C2), 3 kDa (C1B) and 5 kDa (C2) cut-off Vivaspin® 15R concentrators into an NMR buffer. The experiments were carried out on Avance III HD NMR spectrometers (Bruker Biospin), operating at the ¹H Larmor frequencies of 800 MHz (18.8 Tesla) and 600 MHz (14.1 Tesla) equipped with cryogenically cooled probes, and 500 MHz (11.7 Tesla) equipped with a room temperature probe. The temperature was calibrated using deuterated (D4, 98%) methanol for cryogenically cooled probes and protonated methanol for the room temperature probe. Spectra were processed using NMRPipe (Delaglio et al., 1995). The cross-peak intensities were obtained using Sparky (Si et al., 2015). Sequence-specific assignments of the ¹H_N and ¹⁵N resonances for apo C1B-C2 were obtained using ²H-decoupled 3D HN(CA)CB, HNCA(CB), HN(COCA)CB, and HN(CO)CA (Yamazaki et al., 1994) experiments on a [U-¹³C,¹⁵N; 55%-²H] C1B-C2 sample. Resonance assignments for Cd²⁺-substituted C1B (C1B^Cd) were transferred from those for the native Zn²⁺-containing protein (C1B^Zn) and subsequently verified using 3D CBCA(CO)NH and HNCACB (Muhandiram and Kay, 1994) spectra collected at 14.1 Tesla. Resonance assignments for Cd²⁺-bound C1B-C2 were transferred from those for the isolated C1B^Cd and the Cd²⁺-complexed C2 (Morales et al., 2013b) domains. Chemical shift perturbations Δ were calculated between Cd²⁺-free and Cd²⁺-containing C1B-C2 as well as micelle/PDBu bound C1B^Cd and apo C1B^Cd according to the following equation: Δ = Δ δ H 2 + ( 0.152 Δ δ N ) 2 (1) where Δδ_H and Δδ_N are residue-specific ¹H_N and ¹⁵N chemical shift differences. For the NMR-detected binding experiments, the C1B ligand, phorbol-12,13-dibutyrate (PDBu, Sigma-Aldrich) was dissolved in [²H₆] DMSO (Cambridge Isotopes) and added to the sample containing 94 μM of [U-¹⁵N] enriched C1B^Cd in the presence of 10 mM mixed micelles. Mixed micelles comprising [²H₃₈] dodecylphosphocholine, (DPC, Cambridge Isotopes) and 2-dihexanoyl-sn-glycero-3-[phospho-l-serine] (DPS, Avanti Polar Lipids) at a molar ratio of seven to three were prepared as previously described (Stewart et al., 2011). The final concentration of PDBu in the NMR sample was 100 μM.

The four possible Zn/Cd metallated protein states are identified using the following nomenclature: C1B^Zn (native C1B with Zn²⁺ at both structural sites), C1B^Cd (Cd²⁺ at both structural sites), C1B^Zn/Cd (Zn²⁺ at site 1 and Cd²⁺ at site 2), and C1B^Cd/Zn (Cd²⁺ at site 1 and Zn²⁺ at site 2). The fractional populations of those protein species can be defined as: f Z n = I Z n Σ , f Z n / C d = I Z n / C d Σ , f C d / Z n = I C d / Z n Σ , f C d = I C d Σ Σ = I Z n + I Z n / C d + I C d / Z n + I C d (2) where I is the intensity of the corresponding N-H cross peaks in the ¹⁵N-¹H HSQC spectra for H140, I145, and V147. The concentrations of free Cd²⁺ ([Cd²⁺]) and Zn²⁺ ([Zn²⁺]) can be calculated from the following mass balance equations: [ C d 2 + ] 0 P 0 = [ C d 2 + ] P 0 + 2 f C d + f Z n / C d + f C d / Z n (3) 2 = [ Z n 2 + ] P 0 + 2 f Z n + f Z n / C d + f C d / Z n (4) where P₀, [Cd²⁺]₀, and [Zn²⁺]₀ = 2×P₀ are the total concentrations of protein, Cd²⁺, and Zn²⁺, respectively. It is convenient to define the affinities of metal ions to C1B in terms of individual sites. For the single metal-ion bound species, we use the M [n] notation, where M = Zn or Cd, and n = 1 or 2. For example, C1B^Zn[2] defines C1B with site 2 populated by Zn²⁺ and a vacant site 1, and K_a ^Zn[1] defines the association constant for the binding of Zn²⁺ to site 1 when site 2 is already populated by Zn²⁺. The following equilibria describe the binding processes and the associated K_a values: C 1 B Z n [ 2 ] + Z n 2 + ⇌ K a Z n [ 1 ] C 1 B Z n               K a Z n [ 1 ] = [ C 1 B Z n ] [ C 1 B Z n [ 2 ] ] [ Z n 2 + ] (5) C 1 B Z n [ 1 ] + Z n 2 + ⇌ K a Z n [ 2 ] C 1 B Z n         K a Z n [ 2 ] = [ C 1 B Z n ] [ C 1 B Z n [ 1 ] ] [ Z n 2 + ] (6) C 1 B Z n [ 2 ] + C d 2 + ⇌ K a C d [ 1 ] C 1 B C d / Z n         K a C d [ 1 ] = [ C 1 B C d / Z n ] [ C 1 B Z n [ 2 ] ] [ C d 2 + ]   (7) C 1 B Z n [ 1 ] + C d 2 + ⇌ K a C d [ 2 ] C 1 B Z n / C d         K a C d [ 2 ] = [ C 1 B Z n / C d ] [ C 1 B Z n [ 1 ] ] [ C d 2 + ] (8)

The relative affinity of Cd²⁺ and Zn²⁺ to sites 1 and 2 can then be defined as the ratio of the association constants: χ [ 1 ] = K a C d [ 1 ] K a Z n [ 1 ] = [ Z n 2 + ] f C d / Z n [ C d 2 + ] f Z n (9) χ [ 2 ] = K a C d [ 2 ] K a Z n [ 2 ] = [ Z n 2 + ] f Z n / C d [ C d 2 + ] f Z n (10)

The χ[n] (n = 1 or 2) values report on the relative affinities of Cd²⁺ and Zn²⁺ to a given site C1B site when Zn²⁺ populates the other. A similar set of equilibria can be constructed to obtain the relative Cd²⁺ and Zn²⁺ affinities when Cd²⁺ populates the other site: μ [ 1 ] = [ Z n 2 + ] f C d [ C d 2 + ] f Z n / C d (11) μ [ 2 ] = [ Z n 2 + ] f C d [ C d 2 + ] f C d / Z n (12)

χ[n] and μ[n] for sites 1 and 2 (Table 1) were calculated using the NMR cross-peak intensities and the total concentrations of Cd²⁺, C1B, and Zn²⁺ in the system (see Eqs. 1–3). The NMR cross-peaks intensities were determined using the [¹⁵N-¹H] HSQC spectrum of 0.1 mM [U-¹⁵N] C1B^Zn, equilibrated overnight in the presence of 0.1 mM Cd²⁺.

To monitor the kinetics of Cd²⁺ binding to C1B, 2-fold molar excess of Cd²⁺ was added to 200 μM [U-¹³C,¹⁵N; ∼75%-²H] C1B^Zn in 10 mM HEPES buffer at pH 7.2, 75 mM KCl, and 1 mM TCEP. The process of Zn²⁺ replacement with Cd²⁺ was monitored using SOFAST-HMQC NMR experiments that were conducted on a 500 MHz instrument (11.7 Tesla) equipped with a room temperature probe. The first time point started 12 min post Cd²⁺ addition, and each SOFAST HMQC experiment took 15 min. Because the inter-conversion between Zn²⁺- and Cd²⁺-complexed states is in the slow exchange regime, at any time point their fractional population can be determined from the intensities of the corresponding amide cross-peaks in the SOFAST-HMQC spectra. We used the N-H resonances of His102, Cys132, Thr134, Cys135, and Leu150 as the reporters of Cd²⁺ binding to site 1; and Phe114, Cys115, His117, Cys118, Gly119, Ser120, Tyr123, Lys141, and Cys143 as the reporters of Cd²⁺ binding to site 2.