Crystal structures and enzymatic properties of three formyltransferases from archaea: environmental adaptation and evolutionary relationship

Abstract

Formyltransferase catalyzes the reversible formation of formylmethanofuran from N(5)-formyltetrahydromethanopterin and methanofuran, a reaction involved in the C1 metabolism of methanogenic and sulfate-reducing archaea. The crystal structure of the homotetrameric enzyme from Methanopyrus kandleri (growth temperature optimum 98 degrees C) has recently been solved at 1.65 A resolution. We report here the crystal structures of the formyltransferase from Methanosarcina barkeri (growth temperature optimum 37 degrees C) and from Archaeoglobus fulgidus (growth temperature optimum 83 degrees C) at 1.9 A and 2.0 A resolution, respectively. Comparison of the structures of the three enzymes revealed very similar folds. The most striking difference found was the negative surface charge, which was -32 for the M. kandleri enzyme, only -8 for the M. barkeri enzyme, and -11 for the A. fulgidus enzyme. The hydrophobic surface fraction was 50% for the M. kandleri enzyme, 56% for the M. barkeri enzyme, and 57% for the A. fulgidus enzyme. These differences most likely reflect the adaptation of the enzyme to different cytoplasmic concentrations of potassium cyclic 2,3-diphosphoglycerate, which are very high in M. kandleri (>1 M) and relatively low in M. barkeri and A. fulgidus. Formyltransferase is in a monomer/dimer/tetramer equilibrium that is dependent on the salt concentration. Only the dimers and tetramers are active, and only the tetramers are thermostable. The enzyme from M. kandleri is a tetramer, which is active and thermostable only at high concentrations of potassium phosphate (>1 M) or potassium cyclic 2,3-diphosphoglycerate. Conversely, the enzyme from M. barkeri and A. fulgidus already showed these properties, activity and stability, at much lower concentrations of these strong salting-out salts.

Fig. 1.

(A) Phylogenetic relationship of 16S-rRNA sequence from Methanosarcina barkeri, Methanopyrus kandleri, Archaeoglobus fulgidus, Methanothermobacter thermoautotrophicus, Methanothermus fervidus, and Methanococcus jannaschii. (B) Phylogenetic relationship of the formyltransferase amino acid sequence from these euryarchaeota. Tree A was constructed from the data given in Boone et al. (1993); Tree B was calculated from formyltransferase sequences using the Clustal method of the Megaligne program (DNA Star).

Fig. 2.

Sedimentation equilibrium analysis of formyltransferase from Methanosarcina barkeri. (A) Experimental absorbance data A^{1.2 cm}(r) at 280 nm, best fit to the data assuming a monomer/dimer/tetramer model of self-association (—), and calculated local contributions of monomers (---), dimers (- - -), and tetramers (- • - • -). (B) Local difference ΔA^{1.2 cm} at 280 nm between fitted and experimental data. (C) Statistical accuracy of the calculated absorbance contributions of the different oligomers (obtained by integration over the sample volume): changes in the sum of the squared residuals, σ, of fits to the data of A resulting from one nonoptimal absorbance parameter (Schuck 1994). The meaning of the symbols is the same as in A. Initial protein concentration, 0.56 mg/mL; solvent, 0.1 M potassium phosphate at pH 7.2, 0.3 mM DTT; rotor speed, 19,000 rpm; rotor temperature, 4°C.

Fig. 3.

Dependence of the relative amounts of formyltransferase monomers (▪), dimers (□), and tetramers (○) from M. barkeri on the potassium phosphate concentration. Initial protein concentration, 0.35 mg/mL; solvent, potassium phosphate at pH 7.2 at the concentrations indicated, 0.3 mM DTT; rotor speed, 19,000 rpm; rotor temperature, 4°C.

Fig. 4.

Activity of formyltransferase from M. barkeri at 4°C (□) and 37°C (▪) in potassium phosphate at pH 7.2 at the concentrations indicated. The phosphate buffer contained 0.3 mM dithiothreitol. Before starting the reaction, the enzyme (0.4 mg/mL) was incubated at 4°C for 25 h in the same phosphate buffer as used in the respective assay. For assays at 37°C the enzyme solution was diluted 1:20 after incubation. Samples of 2.5 μL were assayed.

Fig. 5.

Structure of the formyltransferase. (A) The tetramer presented as a Ribbon diagram indicates a particularly extended contact region between subunits 1 (red) and 2 (green) and the equivalent subunits 3 (blue) and 4 (orange). (B) The Ribbon diagram of the monomer visualizes the location of the insertion region (blue), the meander region (black circle), and the loop between strands 6 and 7 (black arrow). (C) The stereo C_α-plot of the superimposed monomers of the enzymes from M. barkeri (red), A. fulgidus (yellow), and M. kandleri (green) documents their similar fold, in particular, in the core regions of the two lobes. This figure was generated using the program MOLSCRIPT (Kraulis 1991).

Fig. 5.

Structure of the formyltransferase. (A) The tetramer presented as a Ribbon diagram indicates a particularly extended contact region between subunits 1 (red) and 2 (green) and the equivalent subunits 3 (blue) and 4 (orange). (B) The Ribbon diagram of the monomer visualizes the location of the insertion region (blue), the meander region (black circle), and the loop between strands 6 and 7 (black arrow). (C) The stereo C_α-plot of the superimposed monomers of the enzymes from M. barkeri (red), A. fulgidus (yellow), and M. kandleri (green) documents their similar fold, in particular, in the core regions of the two lobes. This figure was generated using the program MOLSCRIPT (Kraulis 1991).

Fig. 5.

Structure of the formyltransferase. (A) The tetramer presented as a Ribbon diagram indicates a particularly extended contact region between subunits 1 (red) and 2 (green) and the equivalent subunits 3 (blue) and 4 (orange). (B) The Ribbon diagram of the monomer visualizes the location of the insertion region (blue), the meander region (black circle), and the loop between strands 6 and 7 (black arrow). (C) The stereo C_α-plot of the superimposed monomers of the enzymes from M. barkeri (red), A. fulgidus (yellow), and M. kandleri (green) documents their similar fold, in particular, in the core regions of the two lobes. This figure was generated using the program MOLSCRIPT (Kraulis 1991).

Fig. 6.

The electrostatic properties of the formyltransferase tetramer from (A) M. barkeri, (B) A. fulgidus, and (C) M. kandleri. The molecule surface is coated according to the electrostatic potential: The extreme ranges of red and blue represent potentials of −20k_BT and 20k_BT, respectively (where k_B is the Boltzmann constant and T is temperature). The electrostatic surface potential of the enzymes from M. barkeri and A. fulgidus is nearly neutral, and that of the M. kandleri enzyme highly negative, reflecting the dominance of acidic to basic residues. The potentials are calculated under salt-free conditions. The figure was generated using the program GRASP (Nicholls et al. 1993).