Picomole-scale characterization of protein stability and function by quantitative cysteine reactivity

Abstract

The Gibbs free energy difference between native and unfolded states ("stability") is one of the fundamental characteristics of a protein. By exploiting the thermodynamic linkage between ligand binding and stability, interactions of a protein with small molecules, nucleic acids, or other proteins can be detected and quantified. Determination of protein stability can therefore provide a universal monitor of biochemical function. Yet, the use of stability measurements as a functional probe is underutilized, because such experiments traditionally require large amounts of protein and special instrumentation. Here we present the quantitative cysteine reactivity (QCR) technique to determine protein stabilities rapidly and accurately using only picomole quantities of material and readily accessible laboratory equipment. We demonstrate that QCR-derived stabilities can be used to measure ligand binding over a wide range of ligand concentrations and affinities. We anticipate that this technique will have broad applications in high-throughput protein engineering experiments and functional genomics.

Fig. 1.

Representative QCR experiments for ecRBP and SN. (A) SDS-PAGE of time-dependent modification of ecRBP variant L62C with 1 mM IAM-biotin at 47.1 °C, pH 7.6 (left) (labeling times indicated for lanes 2–6). Streptavidin was used to alter the electrophoretic mobility of the labeled protein (streptavidin bands indicated by a, b, and c). Unlabeled fractions were quantified by densitometry and fit with a single exponential to obtain reaction rates (right) at different temperatures (54.6 °C, blue; 51.7 °C, green; 48.9 °C, red; 47.1 °C, orange; 45.2 °C, purple; 44.5 °C, black); corresponding reaction rates are 2.6 × 10^-3, 1.9 × 10^-3, 9.3 × 10^-4, 5.2 × 10^-4, 2.6 × 10^-4, and 2.0 × 10^-4 s^-1, respectively. Error bars represent the estimated uncertainty of the integrated band intensities (∼2%). (B) Labeling of SN variant L36C with IAM-biotin at 35.3 °C, pH 7.6 (left). The (un)labeled forms migrate differently in the gel, enabling ratiometric quantification to obtain reaction rates (right) at different temperatures (38.3 °C, blue; 35.3 °C, green; 32.3 °C, red; 29.3 °C, orange; 26.3 °C, purple; 23.3 °C, black); corresponding reaction rates are 9.2 × 10^-4, 3.6 × 10^-4, 1.2 × 10^-4, 1.5 × 10^-4, 7.2 × 10^-5, and 3.1 × 10^-5 s^-1, respectively. At 29.3 °C, 26.3 °C, and 23.3 °C, k_int and k_label were manipulated by increasing the concentration of IAM-biotin from 1 mM to 3.16 mM.

Fig. 2.

Three factors determine the temperature range at which global unfolding free energies (ΔG_U) can be determined by quantitative cysteine reactivity. The first limits are set by the accuracy of the measurement of the labeling rate constants: an upper limit occurs at a temperature () and free energy (^minΔG_exp) at ∼10 °C above T_m (red-dashed arrows) where the difference of k_label and k_int is within experimental error; a lower limit occurs at a temperature () and free energy (^maxΔG_exp) at ∼10–20 °C below T_m (green-dashed arrows) where increased stability sufficiently reduces k_label (Eqs. 3 and 4) such that it appears to be independent of temperature within experimental error. The second limit is set in some cases where the mechanism of cysteine protection (i.e. local or global unfolding) is dependent on temperature. Such cases manifest themselves as a deviation of the observed temperature dependence of ΔG_U from that expected for global unfolding. It is well established that global unfolding conditions prevail within ∼10–20 °C of T_m (16, 17), which we refer to as the global unfolding window of observation. The black line illustrates a case in which there is no such switch (modeled by Eq. 5) and the GUWO extends over the entire temperature range; the gray line represents switching between global and local unfolding with a concomitant temperature limit for the GUWO (modeled by Eq. 12 of ref. 6). The third limit is set at a point where EX1 conditions prevail and k_close no longer exceeds k_int (not illustrated). This may occur as stability is diminished (ΔG_U < 1 kcal/mol) or if the concentration of thiol probe [P] is too high. Loss of EX2 conditions is manifested as a loss of the linear dependence of k_label on [P] and can be remedied by reducing [P]. The overall temperature range at which observations can be made is the intersection of all three of these conditions (black and gray bars).

Fig. 3.

Temperature dependence of ΔG_U determined by QCR for (A) SN variants F34C (purple) and L36C (black), and (B) ecRBP variants L62C (black) and A188C (purple). Solid lines indicate a fit to a Gibbs-Helmholtz profile (Eq. 5) using a fixed ΔC_p of 3 kcal mol^-1 K^-1. Error bars represent the error of three independent experiments at select temperatures.

Fig. 4.

Ligand concentration dependence of ΔΔG_U for (A) ecRBP variants L62C (purple) at 48.9 °C and A188C (black) at 54.6 °C, and for (B) ecMBP variants T157C (black) and S263C (purple) at 63.3 °C. The solid lines represent the fit of Eq 7 to the data to obtain K_D values. Error bars correspond to the propagated uncertainty of two combined ΔG_U measurements.

Fig. 5.

Effect of ligands and substrate on SN stability. (A) QCR experiments for SN variant L36C in the absence (black) and presence of 1 mM Ca²⁺ (purple) or 50 μM pdTp (orange) at 35.3 °C. Observed rate constants of 3.0 × 10^-4, 1.2 × 10^-4, and 9.2 × 10^-5 s^-1, respectively, correspond to ΔΔG_U values of 0.6 ± 0.2 and 0.7 ± 0.2 kcal mol^-1. (B) Dependence of ΔΔG_U on a 2∶1 molar ratio of Ca²⁺ and pdTp for SN variants F34C (black) and L36C (purple) fit with Eq. 7 to obtain K_D values. (C) QCR experiments at 35.3 °C for SN variant L36C in the absence of substrate (black) and 4.7 μM single-stranded DNA (green), 4.7 μM single-stranded DNA with 1 mM Ca²⁺ (blue), and 12 μM of a 2∶1 molar ratio of Ca²⁺ and pdTp (red). Observed rate constants of 3.0 × 10^-4, 2.8 × 10^-4, 7.5 × 10^-5, and 5.9 × 10^-5 s^-1, respectively, correspond to ΔΔG_U values of 0.1 ± 0.2, 0.9 ± 0.2, and 1.0 ± 0.2 kcal mol^-1. The L36C mutant is enzymatically active (inset; 1% agarose gel): 1.5 kb double-stranded DNA fragment (lane 2) digested completely (lane 1) by incubation with 0.05 μM SN/L36C at 20 °C for 10 min in a buffer of 1 mM Ca²⁺, 25 mM MOPS, 100 mM KCl, and pH 7.6. The k_cat, k_cat/K_m, and K_m of SN for canonical substrate (double-stranded salmon sperm DNA) are ∼90 s^-1, 2 × 10⁶ M^-1 s^-1, and ∼50 μM, respectively, at pH 7 (38).