Microtubule-severing enzymes: From cellular functions to molecular mechanism

Abstract

Microtubule-severing enzymes generate internal breaks in microtubules. They are conserved in eukaryotes from ciliates to mammals, and their function is important in diverse cellular processes ranging from cilia biogenesis to cell division, phototropism, and neurogenesis. Their mutation leads to neurodegenerative and neurodevelopmental disorders in humans. All three known microtubule-severing enzymes, katanin, spastin, and fidgetin, are members of the meiotic subfamily of AAA ATPases that also includes VPS4, which disassembles ESCRTIII polymers. Despite their conservation and importance to cell physiology, the cellular and molecular mechanisms of action of microtubule-severing enzymes are not well understood. Here we review a subset of cellular processes that require microtubule-severing enzymes as well as recent advances in understanding their structure, biophysical mechanism, and regulation.

Figure 1.

Phylogeny of microtubule-severing enzymes. Top: Evolutionary relationship among microtubule-severing AAA ATPases, katanin p60, spastin, and fidgetin, as well as the closely related VPS4 and MSP1 proteins that act on nonmicrotubule substrates. Homologues from animals, plants, and basal eukaryotes were identified with BLASTp and aligned with M-COFFEE (Moretti et al., 2007). Phylogenetic trees were computed with RAxML (Stamatakis, 2006). The black line in the VPS4 AAA domain indicates a β-domain insertion. VPS4 has clear homologues in archaea while microtubule-severing enzymes do not. Bottom: Evolutionary relationship among p80 katanin homologues. Most clades of eukaryotes have a katanin p80 with an N-terminal WD40 domain and a C-terminal con80 domain (conserved p80 domain). WD40-less versions appear to have independently evolved in nematodes and chordates. All human homologues are shown in red.

Figure 2.

Regulation of microtubule array geometry by severing enzymes. (A) Maintenance of a parallel array of cortical microtubules in Arabidopsis by katanin-dependent severing at microtubule crossovers, followed by catastrophe of the new plus end and treadmilling (not shown) and reorientation of the free microtubule. (B) During reorientation of Arabidopsis shoots in response to light, severing at crossovers followed by continued polymerization of new plus ends, and further branched nucleation, result in reorientation of the cortical microtubule array.

Figure 3.

Structural domains in microtubule-severing enzymes. (A) Superposition of the x-ray structure of the AAA domains of spastin (PDB ID 3B9P; Roll-Mecak and Vale, 2008) and katanin (PDB ID 5WC1; Zehr et al., 2017). Spastin AAA domain, gray. Katanin NBD, hot pink; HBD, light green; the N-terminal helix α1 is part of the fishhook and colored cyan; the C-terminal helix α12 specific to AAA ATPases of the meiotic clade is colored orange. (B) Structure of the katanin MIT domain (PDB ID 2RPA; Iwaya et al., 2010). (C) Structure of the complex between the katanin p60 MIT domain and the p80 conserved C-terminal domain (PDB ID 5NBT; Rezabkova et al., 2017); MIT domain, blue; p80 C-terminal domain, yellow. (D) Structure of the spastin MIT domain in complex with CHMP1B (PDB ID 3EAB; Yang et al., 2008). MIT domain, blue; CHMP1B, lavender. For reference, the domain organization in spastin and katanin can be seen in Fig. 1.

Figure 4.

Katanin forms an asymmetric hexamer with a spiral arrangement of the AAA domains. Orthogonal views of the molecular surface of the assembled katanin hexamer in the spiral conformation showing the central AAA ring with radiating arms composed of a poorly conserved flexible linker and an N-terminal MIT domain. The AAA ring structure was determined by single-particle cryo-EM to 4.4 Å (PDB ID 5WC0; Zehr et al., 2017); the MIT domain structure was determined by NMR (PDB ID 2RPA; Iwaya et al., 2010); the structure of the flexible linker (gray) has not been experimentally determined and was modeled to have an approximate span as the one reported from small angle x-ray scattering (Zehr et al., 2017). Protomer P1, green; P2, cyan; P3, blue; P4, orange; P5, purple; and P6, red.

Figure 5.

Structural elements unique to katanin augment the AAA ring and participate in hexamerization. (A and B) Views of the molecular surface of the assembled katanin hexamer in the spiral conformation highlighting important functional elements. Fishhook, cyan; pore loop 1, yellow; pore loop 2, magenta; C-terminal helices, brick red.

Figure 6.

Proposed mechanism for tubulin tail translocation through the katanin central channel. (A) Left: Spiral conformation (PDB ID 5WC0; Zehr et al., 2017) of the hexamer with ATP bound in all six protomers. Right: ATP hydrolysis and release in the boundary protomer P1 transitions the hexamer to the ring conformation (PDB ID 5WCB; Zehr et al., 2017) that closes the gate between P1 and P6, and translocates the pore loops. (B) Schematic of conformational changes triggered by hydrolysis and release of ATP in the boundary protomer P1 that result in closure of the P1–P6 gate and translocation of the pore loops together with the tubulin substrate by ∼20 Å. Adapted from Zehr et al. (2017). Figure is reprinted with permission from Nature Structural and Molecular Biology.