Cystathionine β-Synthase (CBS) Domains Confer Multiple Forms of Mg²⁺-dependent Cooperativity to Family II Pyrophosphatases

Background: Enzymes and transporters may contain regulatory nucleotide-binding cystathionine β-synthase domains.

Results: Both substrate and adenine nucleotides bind cooperatively to four bacterial cystathionine β-synthase domain-containing pyrophosphatases.

Conclusion: Cystathionine β-synthase domains internally inhibit catalysis and impart cooperativity; both effects are modulated by bound nucleotides.

Significance: The observed regulatory complexity may be characteristic of cystathionine β-synthase domain-containing proteins involved in hereditary diseases.

Abstract

Regulated family II pyrophosphatases (CBS-PPases) contain a nucleotide-binding insert comprising a pair of cystathionine β-synthase (CBS) domains, termed a Bateman module. By binding with high affinity to the CBS domains, AMP and ADP usually inhibit the enzyme, whereas ATP activates it. Here, we demonstrate that AMP, ADP, and ATP bind in a positively cooperative manner to CBS-PPases from four bacteria: Desulfitobacterium hafniense, Clostridium novyi, Clostridium perfringens, and Eggerthella lenta. Enzyme interaction with substrate as characterized by the Michaelis constant (K_m) also exhibited positive catalytic cooperativity that decreased in magnitude upon nucleotide binding. The degree of both types of cooperativity increased with increasing concentration of the cofactor Mg²⁺ except for the C. novyi PPase where Mg²⁺ produced the opposite effect on kinetic cooperativity. Further exceptions from these general rules were ADP binding to C. novyi PPase and AMP binding to E. lenta PPase, neither of which had any effect on activity. A genetically engineered deletion variant of D. hafniense PPase lacking the regulatory insert was fully active but differed from the wild-type enzyme in that it was insensitive to nucleotides and bound substrate non-cooperatively and with a smaller K_m value. These results indicate that the regulatory insert acts as an internal inhibitor and confers dual positive cooperativity to CBS domain-containing PPases, making them highly sensitive regulators of the PP_i level in response to the changes in cell energy status that control adenine nucleotide distribution. These regulatory features may be common among other CBS domain-containing proteins.

Introduction

Pyrophosphate (PP_i) is formed as a by-product of nearly 200 biosynthetic reactions in which nucleoside triphosphates are converted to nucleoside monophosphates and PP_i (1). Many of these PP_i-generating reactions are inhibited by PP_i, highlighting the importance of mechanisms that strictly control the free cellular concentration of PP_i. This control is generally achieved by the hydrolysis of PP_i to phosphate via the action of specific enzymes, soluble inorganic pyrophosphatases (PPases⁴; EC 3.6.1.1), belonging to three non-homologous families (2–4).

Of these PPases, family II soluble PPases, found in prokaryotes, are the most active (k_cat ≈ 10⁴ s⁻¹) and structurally diverse. These homodimeric enzymes are Mn²⁺ or Co²⁺ metalloproteins but additionally require Mg²⁺ for catalysis (5), which occurs by direct attack of a hydroxide ion coordinated with three metal ions (6). Each subunit is formed by two domains connected by a flexible linker with the active site located between the domains (7, 8), making the enzyme ideally suited for allosteric regulation. Indeed, a quarter of the nearly 500 putative family II PPase sequences generally contain a ∼250-residue insert comprising a pair of cystathionine β-synthase (CBS) domains (Bateman module (9)), which have regulatory functions in several other proteins (10, 11–14), and a DRTGG domain of unknown function (see Fig. 1). CBS domains are quite common in nature and are found, for instance, in 75 human proteins and eight proteins of Escherichia coli. Mutations in CBS domains of human enzymes and membrane channels are associated with hereditary diseases, including homocystinuria, retinitis pigmentosa, Bartter syndrome, osteopetrosis, and others (13). CBS domains are self-sufficient units (see Fig. 1B) that retain structure after being removed from functional cores. Moreover, some regulatory proteins consist of only a Bateman module (15, 16). The majority of CBS domain-containing proteins form homodimers with the CBS domains participating in subunit interface interactions. Consequently, the two CBS domain pairs form a disklike structure (10). Interestingly, PPase appears to be the only Bateman module-containing protein in which a DRTGG domain intercalates the CBS domains (see Fig. 1). However, in rare cases, for example Moorella thermoacetica PPase (mtPPase), the enzyme contains only CBS domains in the regulatory part of the molecule. Typically, CBS domains bind adenine nucleotides, such as S-adenosylmethionine, AMP, ADP, and ATP, as regulatory ligands (10, 11–14). Each CBS domain contains a potential binding cavity for nucleotides; however, with rare exceptions (16, 17), each pair of CBS domains generally binds only one adenylate ligand. The structural and kinetic basis for regulation through CBS domains is still unknown for any protein, and CBS domain-containing PPases represent a good model enzyme for exploring this general phenomenon.

FIGURE 1.

CBS-PPase structure. A, domain arrangement in the primary structures of wild-type dhPPase and its deletion variant. The five domains that form one subunit are shown in different colors and are labeled; short linker regions are depicted in black. DHH and DHHA2 are catalytic domains. Numbers indicate residues that start/end domains or their parts or precede/follow the deleted sequences. Residue numbering is based on the sequence of the full-length protein. For dhPPaseΔCDC, the linker sequence is shown using single letter notation. B, two views of the modeled three-dimensional structure of the cpCBS-PPase homodimers (19) in a sphere representation. Domain colors of one subunit are the same as in A; the other subunit is colored gray. The AMP molecules bound between CBS domains are shown in green.

We reported previously that mtPPase (18) and the CBS domain-containing PPase of Clostridium perfringens (cpPPase) (19) are highly sensitive to micromolar concentrations of adenine nucleotides. Remarkably, AMP and ADP acted as inhibitors, whereas ATP and AP₄A were activators. We also determined the three-dimensional structures of an isolated regulatory insert of cpPPase comprising two CBS domains and one DRTGG domain in complexes with AMP and AP₄A (19). Rates of AMP-induced conformational transitions were much faster in cpPPase (20) than in mtPPase (21), suggesting a role for the DRTGG domain in regulatory site performance.

Here, we show that the regulatory part of the molecule, comprising two CBS domains with or without an intercalating DRTGG domain, binds adenine nucleotides cooperatively and confers catalytic cooperativity to cpPPase and three newly isolated family II PPases of Desulfitobacterium hafniense (dhPPase), Clostridium novyi (cnPPase), and Eggerthella lenta (elPPase). We found that both types of cooperativity depend on Mg²⁺ concentration and the identity of the bound nucleotide, further expanding our understanding of the mechanisms involved in regulating cellular PP_i level. These findings were corroborated by studies of a genetically engineered dhPPase deletion variant corresponding to the isolated catalytic part of the enzyme. CBS domain-containing PPases are thus the first known oligomeric PPases whose subunits exhibit cooperative behavior.

EXPERIMENTAL PROCEDURES

Cloning

Genomic DNA extracted from D. hafniense, C. novyi, and E. lenta (DSM numbers 10664, 14992, and 2243, respectively) was obtained from DSMZ (Deutsche Sammlung von Microorganismen und Zellkulturen GmbH). The sequences of full-length dhPPase (residues 1–544; YP_002458003), cnPPase (residues 1–548; YP_878001), and elPPase (residues 1–455; YP_003181314) were amplified by PCR and ligated into NdeI and XhoI restriction sites of the pET28bTEV plasmid (Novagen) for dhPPase and pET46 Ek/LIC plasmid (Novagen) for cnPPase and elPPase. A synthetic gene encoding the catalytic part of dhPPase (dhPPaseΔCDC) containing codons for dhPPase fragments 1–63 and 306–544 connected by the linker ESVEAKQK was obtained from GeneArt. The linker sequence was identical to that found in the family II PPase from Desulfosporosinus acidiphilus SJ4 that most closely resembles the catalytic part of dhPPase but has no regulatory element. The gene was subcloned into the pET28bTEV plasmid using NdeI and XhoI restriction sites. The plasmid constructs thus obtained expressed dhPPase, dhPPaseΔCDC, cnPPase, elPPase, or dhCDC with a cleavable His₆ tag at their N termini. All constructs were confirmed by sequencing.

Protein Expression and Purification

Full-length dhPPase and dhPPaseΔCDC were produced in the E. coli strain BL21(DE3)-RIL (Stratagene) by cultivating at 37 °C in Terrific Broth medium supplemented with 30 μg/ml each of kanamycin and chloramphenicol. Gene expression was induced by incubating with 1 mm isopropyl β-thiogalactopyranoside for 3 h. Collected cells were suspended in lysis buffer (50 mm Tris-HCl, 500 mm NaCl, 20 mm imidazole-HCl, pH 7.8) and disrupted with an MSE 100-watt ultrasonic disintegrator. The cell lysate was applied to a 2-ml TALON Superflow gravity column (GE Healthcare), and dhPPase was eluted using a buffer containing 50 mm Tris-HCl, 500 mm NaCl, and 0.2 m imidazole-HCl (pH 7.8). Imidazole was removed from eluate fractions containing dhPPase by passing through a HiPrep 26/10 column (GE Healthcare) pre-equilibrated with buffer consisting of 50 mm Tris-HCl (pH 7.8) and 500 mm NaCl. The His₆ tag was removed by digesting overnight with tobacco etch virus (TEV) protease at 20 °C as described by the manufacturer after which the mixture was passed through a TALON Superflow gravity column to remove His₆-containing peptides from the sample. The flow-through was further purified by size exclusion chromatography on a Superdex 200 26/60 (GE Healthcare) column pre-equilibrated with 50 mm MOPS-KOH (pH 7.2), 150 mm KCl, 0.1 mm CoCl₂, and 20 μm diethylenetriaminepentaacetate. cnPPase and elPPase were expressed and purified similarly except that the His₆ tag was not removed, and the buffer in size exclusion chromatography was 25 mm MOPS-KOH (pH 7.2), 50 mm KCl, and 0.1 mm CoCl₂. cpPPase was expressed and purified as described previously (19). Protein solutions were stored at −40 to −80 °C.

The purity of the isolated proteins was assessed by SDS-PAGE using Phast system 8–25% gradient gels (GE Healthcare). Protein concentrations were determined with a Nanodrop spectrophotometer (Thermo Scientific) using A₂₈₀^0.1% values of 0.478 for dhPPase, 0.419 for dhPPaseΔCDC, 0.548 for cnPPase, 0.426 for cpPPase, and 0.493 for elPPase calculated from the corresponding amino acid compositions with ProtParam. Molar concentrations were calculated on the basis of the subunit molecular masses of 60.4, 34.5, 63.6, 60.8, and 52.5 kDa, respectively.

Kinetic Assays

The activity assay medium contained 140 μm PP_i (yielding 50 μm MgPP_i complex), 5 mm MgCl₂, and 0.1 m Tes-KOH (pH 7.2) except where specified otherwise. In measurements done at higher Mg²⁺ concentrations, buffer concentration was decreased appropriately to maintain constant ionic strength. The reaction was initiated by adding enzyme, and P_i accumulation due to PP_i hydrolysis was continuously recorded for 2–3 min at 25 °C using an automatic P_i analyzer (22). The analyzer utilizes the principle of color change of methyl green in the presence of phosphomolybdate and consists of a three-channel peristaltic pump and a flow-through photometer. The pump delivers a constant flow of sample, acid molybdate, and methyl green solutions, which are successively combined and directed to the flow photometer. The sensitivity of the analyzer was varied to yield full-scale recorder values of 4 or 20 μm P_i using an appropriate choice of pump tubing diameters; corresponding sample consumption rates were 5 and 1 ml/min. The dead time of the analyzer (i.e. the time from feeding the sample into the analyzer to mixing with the acid molybdate solution, which quenches the enzymatic reaction) was 10–20 s in the standard mode. The precision of initial velocity estimates for reactions yielding nonlinear time courses of P_i production was increased by decreasing the dead time to 1 s using an appropriate modification of the inlet system (23). The initial velocity values obtained in replicate measurements agreed within 5–10% and were strictly proportional to enzyme concentration.

Calculations and Data Analysis

The values of the apparent dissociation constants for the various metal ion complexes used to calculate the free metal ion concentrations at pH 7.2 were as follows: MgPP_i, 112 μm (24); Mg₂PP_i, 2.84 mm (24); MgAMP, 10.5 mm (25); MgADP, 0.42 mm (25); and MgATP, 0.034 mm (25). The values for adenine nucleotides were derived from published values of the corresponding pH-independent constants. Nonlinear least square fits were performed using the program Scientist (Micromath).

Initial velocities of PP_i hydrolysis were typically estimated graphically from the slopes of the tangents to the initial portion of hydrolysis time courses recorded with the P_i analyzer. The time courses generated by truncated PPase were markedly nonlinear because of gradual inactivation of the enzyme upon dilution into the reaction medium. In this case, the initial velocities (v) were obtained by fitting the time courses to Equation 1,

where k_d is the rate constant for enzyme inactivation, t is time, and a is the background signal of the instrument.

The dependence of hydrolysis rate on nucleotide concentration ([N]) at fixed substrate concentration was fit to Equation 2,

where A₀ and A_N are activities of free and nucleotide-saturated enzyme, respectively, and K_N1 and K_N2 are the apparent dissociation constants for the enzyme-nucleotide complexes. Equation 2 was derived for Scheme 1, which describes successive binding of nucleotide to two regulatory sites per enzyme molecule. Here, E₂ refers to a mixture of free and substrate-bound dimeric enzyme. The activity of E₂N is intermediate between the activities of E₂ and E₂N₂ (i.e. activities of two subunits respond independently to nucleotide binding). Equation 2 is equally valid if N is an inhibitor or activator.

SCHEME 1.

Modulation of PPase activity by nucleotide binding at a fixed substrate concentration.

For non-cooperative binding, Equation 3 with a single K_N value was used.

Notably, if Equation 2 instead of Equation 3 is applied to a non-cooperative binding reaction, the fitted values of K_N1 and K_N2 should differ 4-fold: K_N1 = K_N/2 and K_N2 = 2K_N (26). Consequently, the ratio 4K_N1/K_N2 can be used to recognize cooperativity: it is greater than 1 for positive cooperativity and less than 1 for negative cooperativity. In terms of generally accepted nomenclature, K_N is a “microscopic” binding constant, and K_N1 and K_N2 are “macroscopic” binding constants (26). The merit of using 4K_N1/K_N2 instead of h to characterize cooperativity is that the former parameter is directly related to the free energy change associated with cooperativity, that is the effect of a ligand binding at one site on the binding affinity of the other site (27).

Alternatively, nucleotide binding data were fit to the Hill equation (Equation 4) to determine the Hill coefficient h.

Cooperative kinetics of substrate hydrolysis was analyzed in terms of Scheme 2, which assumes different Michaelis constants (K_m1 and K_m2) and equal k_cat values for two active sites in the dimer (E₂); [E]₀ is total enzyme concentration. The values of K_m1, K_m2, and k_cat were obtained by fitting Equation 5 to the dependence of hydrolysis rate (v) on substrate (MgPP_i) concentration at fixed Mg²⁺ concentrations.

As was the case for Equation 2, the ratio 4K_m1/K_m2 can be used to recognize catalytic cooperativity: its value is greater than 1 for positive cooperativity and less than 1 for negative cooperativity (26). The applicability of the microscopic/macroscopic constant concept to Michaelis constants results from the formal similarity of the Michaelis-Menten equation to the equilibrium binding equation (28) despite the fact that they are not equilibrium substrate binding constants in a general case.

SCHEME 2.

Substrate binding and hydrolysis at a fixed Mg²⁺ concentration (S = MgPP_i).

The dependence of K_N on Mg²⁺ (M) concentration was fit to Equation 6 derived for Scheme 3 where (K_N)₀ and (K_N)_M are the limiting values of K_N at zero and infinite Mg²⁺ concentrations, and K_M is the metal binding constant. Equations analogous to Equation 6 were used to analyze the Mg²⁺ concentration dependence of K_m.

SCHEME 3.

Modulation of nucleotide binding to CBS-PPase by Mg²⁺.

RESULTS

Production of PPases

The genes for CBS-PPases and the deletion variant of dhPPase (dhPPaseΔCDC; Fig. 1A) were expressed in E. coli, and the proteins were purified with a yield of ∼10 mg of pure protein/liter of medium. Constructs were designed to encode an additional N-terminal MGSSHHHHHHENLYFQGH sequence containing a His₆ tag to facilitate isolation by immobilized metal ion affinity chromatography. TEV protease cleaves this sequence between the Gln and Gly residues, leaving only a dipeptide extension on the protein. His₆ tag removal did not affect PPase activity but diminished the aggregation of full-size dhPPase during purification and concentration procedures. Tag removal was not performed with dhPPaseΔCDC and other PPases as it did not improve their solubility. Inactive aggregates were separated from soluble active proteins during size exclusion chromatography. The final products were at least 95% pure as estimated by SDS-PAGE.

The final purification step (Superdex 200 chromatography) was conducted in the presence of 0.1 mm Co²⁺ to stabilize the enzymes; Co²⁺ was also routinely added to enzyme stock solutions. The stabilizing effect of Co²⁺ is consistent with data showing that “canonical” family II PPases contain one site that tightly binds transition metal ions (Co²⁺ or Mn²⁺) in addition to several more loosely binding sites with broader specificity that are apparently filled with Mg²⁺ in vivo (5, 29, 30). All of these sites have a role in catalysis, but the tightly bound transition metal ion also has a structural role (7, 8) as it does in many other metalloenzymes. The activities of CBS-PPases preincubated with 0.1 mm Co²⁺ were somewhat higher than those incubated with Mn²⁺. Therefore, Co²⁺ was routinely used as the transition metal cofactor in these studies.

Cooperativity of Substrate Hydrolysis by CBS-PPases

The dependence of CBS-PPase activity on substrate concentration in the presence of 5 mm free Mg²⁺ exhibited systematic deviations from simple Michaelis-Menten kinetics as illustrated in Fig. 2 for dhPPase. The Hill coefficient h, derived from this dependence using Equation 4 by setting A₀ to zero, was 1.37 ± 0.02 (Fig. 2), indicating positive catalytic cooperativity. Therefore, the dependence was analyzed using Equation 5 derived for Scheme 2, which assumes different Michaelis constants (K_m1 and K_m2) for two active sites. This analysis markedly increased the quality of the fit compared with the Michaelis-Menten equation (Fig. 2) as indicated by a ∼12-fold lower sum of the squares of residuals. The values of K_m1 and K_m2 were found to be 26 ± 1 and 10 ± 1 μm, respectively; the ratio 4K_m1/K_m2 (equal to 10) was thus much greater than 1, confirming positive catalytic cooperativity. Importantly, product formation curves were linear, and reaction rates were strictly proportional to enzyme concentration in all cases, ruling out the possibility that the deviation from simple kinetics seen in Fig. 2 resulted from a rapid substrate-dependent change in enzyme oligomeric structure upon dilution into the assay medium.

FIGURE 2.

Rate versus substrate concentration profile for dhPPase measured in the presence of 5 mm Mg²⁺. Enzyme concentration in the assay was 0.1–4 nm. The lines represent the best fits for Equation 5 for cooperative kinetics (solid line) and a simple Michaelis-Menten equation (dashed line). The inset shows a Hill plot of the data where V = k_cat[E]₀; both axes are scaled logarithmically.

The Hill coefficient h as well as the ratio 4K_m1/K_m2 for dhPPase were similarly greater than 1 across the entire 0.05–20 mm Mg²⁺ concentration range (Fig. 3) but decreased at low Mg²⁺ concentrations, indicating a decrease in the degree of cooperativity. This transition resulted from changes in K_m1 but not K_m2, which did not vary significantly (Fig. 3). The dependence of K_m1 on Mg²⁺ concentration obeyed Equation 6 with the parameters (K_N)₀ and (K_N)_M replaced by the respective parameters (K_m1)₀ and (K_m1)_M, the Michaelis constants at zero, and infinite Mg²⁺ concentration. The fitted (K_m1)₀, (K_m1)_M, and K_M values were 4.7 μm, 60 μm, and 0.6 mm, respectively (Table 1). The measured value of k_cat for dhPPase showed little dependence on Mg²⁺ concentration (0.05–20 mm) and was in the range of 300–350 s⁻¹.

FIGURE 3.

Dependence of kinetic cooperativity on Mg²⁺ concentration in four CBS-PPases. The panels show (from top to bottom) the catalytic constant k_cat, the Michaelis constants K_m1 and K_m2, and the Hill coefficient h. The enzyme identities are indicated on the top of the panels. Values of k_cat were fit to the equation k_cat = k_cat,0 + (k_cat,M − k_cat0)/{1 + (K_M/[M])ⁿ} (with n equal to 1 and 2 for cnPPase and cpPPase, respectively) where k_cat,0 and k_cat,M are the limiting values of k_cat at zero and infinite Mg²⁺ concentrations, respectively, and K_M is the metal binding constant. The lines for K_m1 and K_m2 show the best fits for Equation 6. The horizontal dotted lines (h = 1) mark the boundary between positive and negative cooperativity. K_m values are measured in terms of the MgPP_i complex. Error bars represent S.E.

TABLE 1.

Kinetic parameters for PP_i hydrolysis derived from the dependence of k_cat, K_m1, and K_m2 on Mg²⁺ concentration

Enzyme	Parameter value
	kcat dependence			Km1 dependence			Km2 dependence
	kcat,0	kcat,M	KM	(Km1)0	(Km1)M	KM	(Km2)0	(Km2)M	KM
	s−1	s−1	mm	μm	μm	mm	μm	μm	mm
dhPPase	NAa	320 ± 20	NA	4.7 ± 0.5	60 ± 20	0.6 ± 0.2	NA	11 ± 1	NA
cnPPase	370 ± 90	820 ± 120	∼2	NA	22 ± 2	NA	24 ± 1	230 ± 80	2.1 ± 0.5
cpPPase	27 ± 5	1310 ± 60	1.05 ± 0.05b	<10	81 ± 3	NDc	80 ± 50	<1	ND
elPPase	NA	70 ± 14	NA	47 ± 2.5	240 ± 10	0.14 ± 0.02	39 ± 3	13 ± 2	2.3 ± 1.3

^a NA, not attendant.

^b Estimated assuming binding of two metal ions with the same binding constant (see Fig. 3 legend for details).

^c ND, not determined.

A similar kinetic analysis was conducted with three other CBS-PPases. The results are summarized in Fig. 3 and Table 1. With cnPPase, K_m1 was virtually independent of Mg²⁺ concentration, whereas K_m2 increased at high [Mg²⁺]. As a consequence, Mg²⁺ exerted an opposite effect on cooperativity compared with dhPPase, and the Hill coefficient dropped to slightly below unity at high [Mg²⁺]. With cpPPase and elPPase, both K_m1 and K_m2 changed with [Mg²⁺] concentration but in opposite directions. Consequently, the degree of cooperativity increased with increasing [Mg²⁺]. The largest effect was observed with cpPPase, which exhibited h values in the range of 1.0–1.8 depending on Mg²⁺ concentration.

The measured value of k_cat was nearly constant over the entire [Mg²⁺] range for elPPase but changed 2-fold for cnPPase and 50-fold for cpPPase (Fig. 3). For the latter enzyme, k_cat depended on the second power of Mg²⁺ concentration (see Fig. 3 legend), indicating that two metal ions are required for the transition from k_cat,0 to k_cat,M.

The above analyses were based on the assumption that the k_cat value for E₂S is half that for E₂S₂ in Scheme 1. Assuming zero activity for the E₂S species increased the sum of the squared residuals by severalfold, indicating significantly worse fits. In contrast, assuming equal activities of E₂S and E₂S₂ decreased the quality of the fits to a lesser extent, making this case a likely alternative. However, if the ratio of the k_cat values for E₂S and E₂S₂ was treated as an adjustable parameter, its best fit value was generally closer to 0.5 than 1.0. Importantly, the values of K_m1, K_m2, and their ratio showed essentially the same trends with the k_cat ratio equal to 0.5 and 1, indicating positive cooperativity, which decreased at low [Mg²⁺] in both cases. We thus conclude that Scheme 1 is the simplest scheme consistent with the data because any value of the k_cat ratio greater than 0.5 implies additional negative catalytic cooperativity because of diminished activity of one of the active sites in the dimeric enzyme.

Kinetic Behavior of dhPPaseΔCDC

Remarkably, dhPPaseΔCDC, which is devoid of the regulatory unit, demonstrated Michaelian kinetics in the presence of 5 mm Mg²⁺ and over the entire 0.1–20 mm Mg²⁺ concentration range as indicated by a 4K_m1/K_m2 value for Scheme 2 and a Hill coefficient close to 1 (Fig. 4 and Table 1). The single microscopic K_m value describing dhPPaseΔCDC kinetics showed little variation in the metal concentration range used, and its values (3.8–5.0 μm) were markedly less than √K_m1K_m2 (the geometric mean of the two K_m values) values for dhPPase (Fig. 4).

FIGURE 4.

Lack of kinetic cooperativity in dhPPaseΔCDC. Left, rate versus substrate concentration profile measured in the presence of 5 mm Mg²⁺. Enzyme concentration in the assay was 0.1–4 nm. The line represents the best fit for a Michaelis-Menten equation. The inset shows a Hill plot of the data; both axes are scaled logarithmically. Right, Mg²⁺ concentration dependence of kinetic parameters. The panels show (from top to bottom) the Michaelis constant K_m1, the ratio 4K_m1/K_m2, and the Hill coefficient h. The horizontal dotted lines (4K_m1/K_m2 = 1 and h = 1) mark the boundary between positive and negative cooperativity. Error bars represent S.E.

The value of k_cat for dhPPaseΔCDC was 100–120 s⁻¹ over the 0.1–20 mm Mg²⁺ concentration range. The somewhat lower activity of dhPPaseΔCDC compared with dhPPase may indicate that deletion of the regulatory part of the protein caused some distortions in the active site.

Cooperativity of Nucleotide Binding to dhPPase

Fig. 5 shows the concentration dependence of nucleotide effects on activity measured at fixed substrate (MgPP_i) and Mg²⁺ concentrations. Micromolar concentrations of AMP and ADP caused inhibition in most cases, whereas ATP acted as an activator. Qualitatively similar effects of these nucleotides were reported previously for the CBS domain-containing mtPPase (18). The major new finding here is that, in most cases where the size of the effect allowed quantitative analysis, the curves were poorly described in terms of a simple 1:1 binding model (Equation 3) but obeyed Equation 2 for cooperative binding to two sites with concomitant activation or inhibition. The quality of the fit was relatively insensitive to the value of A_1/2 in Scheme 1, but better fits were generally observed with A_1/2 = (A₀ + A_N)/2 compared with the limiting cases of A_1/2 = 0 or A_N. Importantly, most curves generated K_N1 values greater than K_N2 regardless of the A_1/2 value used and Hill coefficients greater than 1 (Table 2). Thus, nucleotide binding also exhibited positive cooperativity. In only two pairs (cpPPase/ADP and elPPase/ATP) did the analysis indicate insignificant cooperativity.

FIGURE 5.

Concentration dependence of adenine nucleotide effects on the activities of four CBS-PPases measured at fixed concentrations of substrate (50 μm MgPP_i) and Mg²⁺ (5 mm). The lines show the best fits to Equation 2 with A_1/2 = (A₀ + A_N)/2. Activities without nucleotides (220, 350, 800, and 35 s⁻¹ for dhPPase, cnPPase, cpPPase, and elPPase, respectively) were taken as unity.

TABLE 2.

Parameters describing nucleotide effects on four CBS-PPases as derived from Figs. 5 and 6 and similar data measured at different substrate concentrations

Nucleotide	[MgPPi]	vN/v0a	KN1	KN2	4KN1/KN2	√KN1KN2	h
	μm		μm	μm		μm
dhPPase
AMP	5	0.13 ± 0.02	3 ± 2	0.3 ± 0.2	40 ± 30	0.95 ± 0.05	1.75 ± 0.14
AMP	50	0.037 ± 0.001	4.8 ± 0.7	0.41 ± 0.04	48 ± 10	1.39 ± 0.04	1.68 ± 0.04
AMP	400	0.035 ± 0.006	6.7 ± 1.5	0.5 ± 0.1	53 ± 13	1.83 ± 0.05	1.74 ± 0.05
ADP	5	0.23 ± 0.01	0.38 ± 0.05	0.14 ± 0.02	11 ± 3	0.24 ± 0.01	1.51 ± 0.05
ADP	50	0.09 ± 0.01	0.99 ± 0.09	0.90 ± 0.08	4.4 ± 0.8	0.95 ± 0.02	1.34 ± 0.03
ADP	400	0.05 ± 0.01	1.4 ± 0.1	2.1 ± 0.2	2.6 ± 0.4	1.72 ± 0.04	1.23 ± 0.04
ATP	5	3.2 ± 0.1	2.0 ± 0.3	0.63 ± 0.08	13 ± 3	1.14 ± 0.02	1.55 ± 0.04
ATP	50	2.5 ± 0.2	2.3 ± 0.8	1.5 ± 0.1	6 ± 2	1.9 ± 0.2	1.45 ± 0.13
ATP	400	2.1 ± 0.1	4 ± 1	1.7 ± 0.5	9 ± 5	2.5 ± 0.1	1.48 ± 0.10
cnPPase
AMP	50	0.63 ± 0.01	>2000	<80	>100	390 ± 20	2.3 ± 0.2
ADP	50	1.00	NAb	NA	NA	NA	NA
ATP	50	1.32 ± 0.02	NDc	ND	ND	0.92 ± 0.07	ND
ADPd	50	0.98 ± 0.02	ND	ND	ND	14 ± 1	ND
cpPPase
AMP	50	0.035 ± 0.015	12 ± 5	1.1 ± 0.5	40 ± 20	3.7 ± 0.2	1.7 ± 0.1
ADP	50	0.50 ± 0.02	3.2 ± 0.4	18 ± 3	0.7 ± 0.2	7.7 ± 0.6	0.92 ± 0.06
ATP	50	1.26 ± 0.02	>70	<3	>100	14 ± 2	2.2 ± 0.5
elPPase
AMP	50	1.00	NA	NA	NA	NA	NA
ADP	50	0.32 ± 0.01	11.1 ± 1.6	24 ± 4	1.85 ± 0.6	16 ± 1	1.14 ± 0.07
ATP	50	5.2 ± 0.3	123 ± 14	380 ± 70	1.3 ± 0.4	220 ± 40	1.07 ± 0.07
AMPe	50	1.00f	1100 ± 200	1700 ± 300	2.3 ± 0.7	1350 ± 50	1.22 ± 0.06
ADPg	50	0.17 ± 0.01	3.9 ± 0.5	24 ± 3	0.65 ± 0.15	10 ± 1	0.92 ± 0.05
AMPh	50	0.24 ± 0.01	>250	<10	>100	52 ± 3	2.2 ± 0.2

^a Values of v_N and v₀ are activities extrapolated to infinite concentration of the variable nucleotide and measured in the absence of any nucleotide, respectively.

^b NA, not attendant.

^c ND, not determined.

^d Measured in the presence of 10 μm ATP.

^e Measured in the presence of 500 μm ATP.

^f Manually fixed parameter value.

^g Measured in the presence of 100 μm AMP.

^h Measured in the presence of 20 μm ADP.

Interestingly, ADP did not affect cnPPase activity, and AMP was equally ineffective with elPPase (Fig. 5). Activity was nucleotide-insensitive in these enzyme-nucleotide pairs over the entire 0.05–20 mm Mg²⁺ concentration range.⁵ To test whether the nucleotides bound to enzyme in these cases, we performed competition experiments (Fig. 6A). In both cases, a “functionally silent” nucleotide modulated the activating effect of ATP, indicating that the former is able to bind. Moreover, although ineffective by itself, AMP synergistically enhanced the effect of ADP on elPPase, shifting the inhibition profile to lower ADP concentrations and decreasing the activity value extrapolated to infinite ADP concentration (Fig. 6B). This profile shift is explained by a 3-fold lower K_N1 value for ADP in the presence of 100 μm AMP (Table 2). Accordingly, AMP acted in a concentration-dependent manner to potentiate the inhibitory action of ADP measured at a constant ADP concentration (Fig. 6C). Clearly, binding of AMP to one site greatly enhanced ADP binding to the other site(s) to form a mixed enzyme-AMP-ADP complex with low activity in these conditions. The values in the last line of Table 2 suggest that AMP binding to elPPase is highly cooperative: the first nucleotide molecule binds quite weakly, but the mean binding constant (√K_N1K_N2) is comparable with that of the other nucleotides. It is unlikely that the extraordinarily high values of 4K_N1/K_N2 and h for AMP, which are close to their limiting values for two-site binding (27), are somehow associated with the ADP present in the system because the latter nucleotide exhibited quite low binding cooperativity with elPPase (Table 2).

FIGURE 6.

Functionally silent nucleotides as modulators of the effects of “functionally active” nucleotides in CBS-PPases. A, ADP reversal of ATP activation of cnPPase and AMP reversal of ATP activation of elPPase. The fixed ATP concentration was 500 and 10 μm in experiments with elPPase and cnPPase, respectively. B, ADP inhibition of elPPase in the presence of 100 μm AMP. The dependence without AMP (dashed line) was taken from Fig. 5. C, AMP inhibition of elPPase in the presence of 20 μm ADP. The reaction medium contained 50 μm MgPP_i and 5 mm Mg²⁺ in all cases. Activities without any nucleotide were taken as 100%. The lines show the best fits to Equation 2. rel., relative.

The above analysis of nucleotide effects was conducted at a half-saturating substrate concentration. Because the substrate itself also binds cooperatively, the apparent nucleotide binding cooperativity may result from nucleotide effects on substrate binding. Therefore, measurements were extended to low and high substrate concentrations at which the enzyme (dhPPase) predominantly exists in the substrate-free form or as enzyme-substrate complex, respectively. As Table 2 makes clear, the cooperativity of AMP and ATP binding as characterized by 4K_N1/K_N2 and h did not depend significantly on substrate concentration and showed only a moderate change in the case of ADP binding. Therefore, the observed cooperativity of nucleotide binding is not induced by bound substrate or by modulation of substrate binding. In contrast, the relative final activity at infinite nucleotide concentration (v_N/v₀) increased with increasing substrate concentration in all cases, whereas the mean binding affinity (√K_N1K_N2) significantly decreased.

The Mg²⁺ dependence of AMP, ADP, and ATP binding to dhPPase and of AMP binding to cpPPase was determined (Fig. 7). These enzymes and nucleotides were chosen because they demonstrate profound effects on activity, allowing accurate analysis of the binding curves. Although quite large, the effect of ATP on elPPase was not included in this analysis because ATP chelates Mg²⁺ appreciably (25) at the high ATP concentrations at which this enzyme operates (Fig. 5), introducing uncertainty in free Mg²⁺ concentration at the lower values in this range. Varying Mg²⁺ concentration in the assay changed K_N1 and K_N2 values for AMP binding to dhPPase and cpPPase in opposite directions (Fig. 7). As a result, the ratio 4K_N1/K_N2, the Hill coefficient h, and hence the degree of cooperativity increased with increasing Mg²⁺ concentration. With ATP, K_N1, but not K_N2, was Mg²⁺-dependent, resulting in a similar although smaller effect. With ADP, both binding constants changed in parallel without affecting 4K_N1/K_N2 or h. Notably, dhPPase activity at saturating ATP concentration (470 ± 10 s⁻¹) was independent of [Mg²⁺].

FIGURE 7.

Mg²⁺ concentration dependence of nucleotide binding cooperativity in dhPPase and cpPPase. The panels show (from top to bottom) K_N1 (○) and K_N2 (●) values, the ratio 4K_N1/K_N2, and the Hill coefficients. The enzyme and nucleotide identities are indicated on the top of the panels. The K_N1 and K_N2 lines show the best fits for Equation 6. The horizontal dotted lines (4K_N1/K_N2 = 1 and h = 1) mark the boundary between positive and negative cooperativity. Error bars represent S.E.

The dependences of K_N1 and K_N2 on Mg²⁺ concentration ([M]) obeyed Equation 6 derived for a simple model that assumes that metal binding to a site characterized by the dissociation constant K_M shifts the dissociation constant for the enzyme-nucleotide complex from (K_N)₀ to (K_N)_M. The fitted values of these parameters are summarized in Table 3. Interestingly, K_M values were essentially independent of the nucleotide and enzyme used and correlated well with the K_M value derived from substrate hydrolysis kinetics for dhPPase (and other PPases) (Table 1). Moreover, in all cases, the K_M values derived from K_N2 dependence were greater than those derived from the K_N1 dependence; in both cases, their ratio was not much different from 4, the value expected for non-cooperative metal binding to two identical sites.

TABLE 3.

Kinetic parameters for nucleotide inhibition derived from the dependencies of K_N1 and K_N2 on Mg²⁺ concentration (Fig. 7)

Substrate (MgPP_i) concentration was fixed at 50 μm.

Nucleotide	Parameter value
	KN1			KN2
	KN1,0	KN1,M	KM	KN2,0	KN2,M	KM
	μm	μm	mm	μm	μm	mm
dhPPase
AMP	1.7 ± 0.1	190 ± 100	0.6 ± 0.1	7 ± 3	0.9 ± 0.3	1.4 ± 0.9
ADP	0.73 ± 0.03	1.73 ± 0.06	0.5 ± 0.1	1.02 ± 0.04	3.2 ± 0.2	1.3 ± 0.2
ATP	0.83 ± 0.07	2.9 ± 0.2	0.3 ± 0.1	NAa	NA	NA
cpPPase
AMP	0.6 ± 0.1	26 ± 11	0.3 ± 0.1	5.6 ± 0.2	2.4 ± 0.2	1.6 ± 0.5

^a NA, not attendant.

Effects of Nucleotides on Kinetic Cooperativity

Table 4 summarizes nucleotide effects on hydrolysis kinetics measured at a Mg²⁺ concentration of 5 mm. The general trend was that nucleotides decreased kinetic cooperativity as characterized by h values. With cnPPase, which exhibited no cooperativity at this Mg²⁺ concentration (Fig. 3), ATP conferred slightly negative cooperativity (h = 0.86 ± 0.02). Another general trend was that the activating nucleotide (ATP) increased k_cat values up to 2-fold and decreased the mean K_m value (√K_m1K_m2) up to 3-fold for all CBS-PPases except cpPPase in which the latter parameter increased in the presence of ATP. This means that ATP inhibits rather than activates cpPPase at low substrate concentrations.

TABLE 4.

Nucleotide effects on the kinetic parameters for PP_i hydrolysis in the presence of 5 mm Mg²⁺ estimated using Equation 5

Nucleotide	kcat	Km1	Km2	√Km1Km2	h
	s−1	μm	μm	μm
dhPPase
None	350 ± 20	26 ± 1	10 ± 1	20 ± 1	1.29 ± 0.02
ATP (10 μm)	450 ± 10	10 ± 1	11 ± 1	10 ± 1	1.24 ± 0.02
cnPPase
None	540 ± 40	23 ± 2	80 ± 30	44 ± 3	1.02 ± 0.08
AMP (2 mm)	400 ± 10	25 ± 1	65 ± 4	42 ± 1	1.07 ± 0.01
ADP (1 mm)	540 ± 30	20 ± 1	80 ± 20	44 ± 5	0.95 ± 0.04
ATP (100 μm)	670 ± 10	5.1 ± 0.1	45 ± 3	15 ± 1	0.86 ± 0.02
cpPPase
None	1080 ± 30	80 ± 30	4 ± 2	18 ± 1	1.8 ± 0.1
ADP (500 μm)	1080 ± 40	122 ± 19	28 ± 7	60 ± 7	1.50 ± 0.09
ATP (500 μm)	1700 ± 40	63 ± 5	32 ± 5	47 ± 4	1.35 ± 0.06
elPPase
None	89 ± 4	220 ± 70	18 ± 8	63 ± 3	1.7 ± 0.1
AMP (1 mm)	88 ± 4	170 ± 20	39 ± 9	87 ± 10	1.47 ± 0.08
ADP (500 μm)	21 ± 1	7.0 ± 0.3	32 ± 5	15 ± 1	0.98 ± 0.03
ATP (1 mm)	180 ± 10	38 ± 4	32 ± 7	36 ± 4	1.27 ± 0.08

Of the three inhibition cases documented in Table 4, one (cnPPase/AMP) resulted solely from a decrease in k_cat (noncompetitive inhibition), and one (cpPPase/ADP) resulted from an increase in √K_m1K_m2 (competitive inhibition). In the last case (elPPase/ADP), the inhibitor decreased both k_cat and √K_m1K_m2 to the same degree (uncompetitive inhibition), indicating an absence of inhibition at very low substrate concentrations. Functionally silent nucleotides (ADP with cnPPase and AMP with elPPase; see Fig. 5) did not significantly change k_cat or √K_m1K_m2 (Table 4).

Similar analyses could not be performed in the cases of the most efficient inhibitors (AMP and ADP with dhPPase and AMP with cpPPase). These pairs exhibited quite low (although finite) activities of the enzyme-nucleotide complexes, resulting in a significant relative contribution of nucleotide-free enzyme to activity even at high nucleotide concentrations.

CBS-PPase Activity as a Function of ADP:ATP Ratio

The ADP:ATP ratio is a critical determinant of cellular energy status that regulates many metabolic activities. Fig. 8 shows how the value of this parameter affected activities of the four CBS-PPases. The sum of ADP and ATP concentrations was fixed at 0.5 mm. Although this value is an order of magnitude less than that in a typical cell, this did not markedly affect the profiles because ADP and ATP concentrations exceeded the respective √K_N1K_N2 values (Table 2) in most cases. Substrate concentration was arbitrarily fixed at 50 μm. Because ATP is an activator and ADP is an inhibitor or an inert species, enzyme activity decreased with increasing ADP:ATP ratios in all cases. However, the sensitivity of CBS-PPases to this parameter varied considerably with dhPPase being the most sensitive and cnPPase being the least sensitive.

FIGURE 8.

Activities of CBS-PPases as a function of [ADP]:[ATP] ratio. The reaction medium contained 50 μm MgPP_i, 5 mm Mg²⁺, and different concentrations of ADP and ATP such that their sum was 0.5 mm. Activity in the presence of 0.5 mm ATP was taken as unity in all cases. Error bars represent S.D.

DISCUSSION

The organisms that harbor the CBS-PPases used in this study are Gram-positive anaerobic bacteria belonging to the phylum Firmicutes (D. hafniense, C. novyi, C. perfringens) or Actinobacteria (E. lenta). Three of them (C. novyi, C. perfringens, and E. lenta) are pathogenic and are found in the human intestinal tract. D. hafniense is found in soil and is capable of reductive dechlorination of chlorophenols (31). In three of these bacteria (excluding C. perfringens), PP_i metabolism depends on soluble and membrane-bound PPases (32), which convert metabolically inert and inhibitory PP_i into usable P_i. The soluble PPase converts PP_i energy into heat. In contrast, membrane-bound PPases use PP_i to establish an H⁺ gradient (D. hafniense) or Na⁺ gradient (C. novyi and E. lenta) and thereby mobilize PP_i energy to maintain cation gradients and help cells survive under stress conditions (33). This dual utilization of PP_i explains the need for regulation of soluble PPases, which has been achieved evolutionarily by the acquisition of a pair of regulatory, energy-sensing CBS domains. Binding of ATP, which prevails under high energy conditions, activates soluble PPases and decreases PP_i levels, thereby facilitating biosynthetic reactions that are inhibited by PP_i. Under low energy conditions, bound ATP is replaced by ADP and AMP, resulting in inhibition of PPase and an increase in the PP_i level, which in turn slows down biosynthesis and provides substrate for membrane PPases that support the membrane H⁺ or Na⁺ gradient.

The results reported above reveal added complexity in PPase regulation: the CBS domains introduce dual positive cooperativity to the regulatory mechanism. The first level of cooperativity relates to nucleotide binding and serves to amplify the initial signal (change in adenine nucleotide distribution). Simple calculations for AMP inhibition of dhPPase (Fig. 5) with an h value of 1.68 (Table 2) showed that a change in PPase activity from 10 to 90% of its maximal value occurs when AMP concentration changes 16-fold, whereas the same effect would require an 81-fold change in AMP level if it bound non-cooperatively. Cooperativity of nucleotide binding is not caused by bound substrate as its degree, characterized by the values of 4K_N1/K_N2 and h in Table 2, does not depend (or only slightly depends) on the degree of enzyme saturation with substrate. Notably, nucleotide binding cooperativity is not uncommon among other CBS domain-containing proteins. Positive cooperativity was reported for S-adenosylmethionine binding to cystathionine β-synthase (34), AMP binding to IMP dehydrogenase (13), and AMP and ATP binding to AMP-activated protein kinase (13). More recently, AMP-activated protein kinase was reported to successively bind nucleotides with decreasing affinity (17, 35), which suggests negative binding cooperativity.

The second level of positive cooperativity in CBS-PPases relates to enzyme interaction with substrate as characterized by the Michaelis constant (K_m). This type of behavior is called “catalytic cooperativity” to distinguish it from the more common “binding cooperativity.” One should keep in mind that catalytic cooperativity may not be interpreted in terms of binding insofar as K_m is not a thermodynamic binding constant in a general case and is a combination of six individual rate constants in the case of PPase (36). Therefore, a change in K_m value may result from a change in any of the rate constants that describe the four catalytic steps in PPase: substrate binding, substrate hydrolysis, and sequential release of two phosphates.

Catalytic cooperativity, which further amplifies the initial signal, is observed in the absence of nucleotides, is modulated by nucleotides (Table 4), and is therefore induced by the regulatory insert itself rather than by the bound nucleotide. Consistent with this notion, removal of the insert abolished the cooperativity. The capacity of CBS domains to impart cooperativity on enzyme or transporter function has not been previously recognized. However, Labesse et al. (37) have recently reported that IMP dehydrogenase binds its substrate, IMP, with positive cooperativity; this cooperativity was abolished by mutation of a single residue in the CBS domain or binding of MgATP to the enzyme. Of note, CBS-PPases are the only cooperative enzymes among different types of oligomeric PPase enzymes.

Mg²⁺ binding had a strong modulatory effect on both types of cooperativity, but its mode of action varied with enzyme and nucleotide. Increasing Mg²⁺ concentration generally conferred or increased positive kinetic and nucleotide binding cooperativity (Figs. 3 and 7). However, Mg²⁺ changed the kinetic cooperativity in cnPPase in the opposite direction (Fig. 3) and did not affect ADP binding cooperativity in dhPPase (Fig. 7). Similar values of the Mg²⁺ binding constant, K_M, derived from measurements of hydrolysis kinetics (Table 1) and nucleotide effects (Table 3) suggest that a single site controls both cooperativity types. Notably, the structure of the AMP-bound regulatory portion of cpPPase crystallized in the presence of Mg²⁺ contains no bound divalent metal ion (19). In contrast, canonical family II PPases, which represent the catalytic part of CBS-PPases, bind three or four Mg²⁺ ions in the absence of substrate (5) and up to five metal ions in its presence (6). All metal ions are located in the active site, but one of them is not absolutely required for activity (6). This last activity-modulating site is thus the most likely candidate for the site that modulates cooperativity in CBS-PPases.

All K_N values are reported here in terms of total nucleotide concentrations without regard to complexation with Mg²⁺. Based on the Mg²⁺ binding constants (see “Experimental Procedures”), 99.8% of ATP, 98% of ADP, and 70% of AMP exist as magnesium complexes at the highest Mg²⁺ concentration used (20 mm). Thus, it is not unlikely that nucleotide-bound Mg²⁺ modulates enzyme-nucleotide interactions and therefore contributes to the dependences shown in Fig. 7. However, several lines of evidence rule out the possibility that Mg²⁺ binding to nucleotides is the sole cause of the Mg²⁺ effects on K_N values. As already mentioned, two molecules of AMP bind to cpPPase as free nucleotide (19). If recalculated in terms of free AMP, K_N1 and K_N2 values for dhPPase and cpPPase (Fig. 7) would decrease in parallel (up to 3-fold at the highest Mg²⁺ concentration), resulting in a minor effect on the AMP binding pattern. The crystal structures of the cpPPase fragment (19) suggest that other nucleotides form major contacts with the enzyme through their AMP moieties. Therefore, free and Mg²⁺-complexed ADP or ATP are likely to have equal or similar affinities for enzyme. This interpretation is consistent with similar mean K_N values (√K_N1K_N2) for AMP, ADP, and ATP observed for dhPPase and cpPPase, which bind all three nucleotides. However, the possibility that Mg²⁺ binding to ADP and ATP contributes to their binding patterns in Fig. 7 cannot be excluded.

The structural basis for the effects of the regulatory element on the active site and for the modulation of these effects by bound nucleotides still remains to be determined for PPase as well as for other CBS domain-containing enzymes and transporters. Our results pose further questions in this context. Because the two AMP molecules bound per dimeric regulatory fragment of cpCBS-PPase are in proximity to each other (19), one would expect negative rather than positive cooperativity in their binding. Indeed, AMP analog binding was shown to exert negative cooperativity in mtPPase (21). The reversal of nucleotide binding cooperativity in PPase observed in the present study may reflect plasticity of CBS domains and is consistent with the known variability of their amino acid sequences (10). The available crystallographic data show significant opening of the CBS domain interface in the activator (AP₄A) complex of cpPPase compared with that in the inhibitor (AMP) complex (19). Lucas et al. (38) observed an open-to-closed conformational change in the CBS domain pair of Methanocaldococcus jannaschii protein MJ0100 upon successive binding of adenosine derivatives. Quite recently, Jeong et al. (39) reported a dramatic AMP-induced structural change in Arabidopsis thaliana protein CBSX2.

The concept of “autoinhibition” was introduced vis-à-vis CBS domains by Janosík et al. (40) to explain cystathionine β-synthase activation by S-adenosyl-l-methionine as well as its limited proteolysis and thermal denaturation. According to this concept, the CBS domain insert acts as an “internal inhibitor,” sensitizing proteins with catalytic or transport functions to structural changes caused by nucleotide binding and permitting either activation or further inhibition. CBS domain-containing PPases provide strong support for this theory. Thus, deletion of the regulatory domains in dhPPase favorably changed its K_m1 value (Table 3). A similar deletion in cystathionine β-synthase increased k_cat but not K_m (41, 42). The concept of autoinhibition is also consistent with the diminished activity (1–3 orders of magnitude) of CBS domain-containing PPases compared with their close homologs lacking CBS domains (5) as well as their ability to be activated by ATP. Many mutations in the CBS domains also activate mtPPase, and some even reverse the effects of nucleotide from inhibition to activation (43), resembling the effects of mutations in cystathionine β-synthase (44) and AMP-dependent protein kinase (45, 46).

Unlike other known CBS domain-containing proteins, dhPPase, cnPPase, and cpPPase contain a DRTGG domain insert within the CBS domain pair. The results of the present work do not shed light on the function of this extra domain in these (and most other) CBS-PPases as we did not observe any fundamental differences in the behavior of these PPases compared with that of elPPase, which has no DRTGG domain in its structure.

In summary, CBS-PPase presents a unique case, demonstrating in one protein the diverse regulatory capabilities of CBS domains as sensors of cell energy status. The regulation of this enzyme is based on a complex interplay among the effects of nucleotides, substrate, and Mg²⁺ on binding cooperativity. Further work is clearly needed to fully elucidate this interplay and determine its importance for other CBS domain proteins.