6.4 The chitin component

Chitin synthesis is the responsibility of the enzyme chitin synthase, an integral membrane enzyme that catalyses the transfer of N-acetylglucosamine from uridine diphosphate (UDP)-N-acetylglucosamine to a growing chitin chain (described briefly in section 5.15 Subcellular components of eukaryotic cells: fungal cell wall; CLICK the title to view the page). Most fungi have several chitin synthase genes, which suggests some overlap and/or specialisation in function. However, the significance of each of the enzymes in the family is not known. The classic approach is to find or induce mutations to determine the effect of the defective enzyme(s), but mutations in common chitin synthase genes do not always result in similar phenotypes. Paradoxically, among the mutants identified are those with reduced chitin content in the walls, but normal chitin synthase activity in in vitro assays, and a second type which has drastically reduced enzyme activity in in vitro assays but has the usual cell wall chitin content. In addition, other genes, also named CHS, are not associated with chitin synthase enzyme activity but are involved in the regulation or localisation of that activity. Some chitin synthases are zymogens that are activated by proteolysis, and others are regulated by phosphorylation. Some fungi have more than 20 CHS genes, and some have only 1 (see Gow et al. (2017) for references).

The multiple chitin synthase (CHS) genes have been grouped into two families and seven classes, based on amino acid sequence (see Section 5.15). Class I, II and IV genes are present in all fungi; classes III, V, VI and VII are specific to filamentous fungi and some dimorphic species. Chitin synthases of classes I, II and III all share a catalytic domain surrounded by a hydrophilic aminoterminal region and a hydrophobic carboxyterminal region. The number of chitin synthase genes within each species varies and the exact functions of many chitin synthases are still unknown (Rogg et al., 2012). In some fungi the chitin synthase activity is fused to their molecular motor (Section 5.13); both domains being required for correct cellular function. Class V and class VII chitin synthases, which are exclusive to filamentous fungi, encode proteins of around 1,500 amino acids that contain a myosin motor-like domain in their aminoterminal region, which has been shown to interact directly with the actin cytoskeleton (Steinberg, 2011; Fernandes et al., 2016; Morozov & Likhoshway, 2016). Examples of these myosin motor/chitin synthase fusion proteins have been found in Aspergillus nidulans, and in fungi pathogenic to humans (Coccidioides posadasii and Paracoccidioides brasiliensis), and plant pathogens too (Colletotrichum graminicola and Pyricularia oryzae). These enzymes are often essential for growth, morphogenesis, virulence, and stress tolerance. The motor domain functions in actin-mediated cytoplasmic transport to the hyphal apex. However, the motor domain is not required for vesicle motility in Aspergillus nidulans and Ustilago maydis, and the motor may instead serve to tether vesicles at the hyphal apex (Treitschke et al., 2010), increasing the residence time at its location and favouring vesicle fusion with the plasma membrane (Schuster et al., 2012). Interestingly, in Ustilago maydis the class VII chitin synthase Mcs1 and the class V chitin synthase 6 (Chs6) can be cotransported on the same secretory vesicle along with the β1,3-glucan synthase Gsc1. The complex of glucan and chitin synthases seems to be tethered to a localised site of exocytosis and wall synthesis (Schuster et al., 2016).

The different chitin synthase isozymes can be associated with a variety of specific functions that are time or position dependent, such as formation of septa and spores in Aspergillus (Ichinomiya et al., 2005), appressoria in Colletotrichum (Werner et al., 2007) and complexing with chitin deacetylases to produce chitosan in Cryptococcus (Banks et al., 2005), or for wall repair or response to stress (Bowman & Free, 2006). Some can even be shown to be essential for pathogenicity in several species, in the sense that deletion mutants are non-pathogenic a feature that confirms the critical importance of cell wall integrity in fungal infection processes (Werner et al., 2007; Martín-Urdíoz et al., 2008).
But the most essential function to which the chitin synthases contribute is hyphal extension growth and we are beginning to understand how these proteins are localised in the cell and how they are transported to their sites of action in regions of active wall growth. As described in Chapter 5, hyphae of septate filamentous fungi have a unique organelle, the Spitzenkörper, which is an accumulation of vesicles, ribosomes and cytoskeletal components at their growing apex (CLICK HERE to view the page in Chapter 5). It is also clear that fungal cells have two well-defined types of secretory vesicle: macrovesicles are conventional eukaryotic secretory vesicles, which carry the components of the cell wall matrix, glucans, extracellular enzymes and glycoproteins; and these are accompanied by a large population of microvesicles carrying chitin synthase. These microvesicles have been called chitosomes; they are the smallest vesicles that have the ability to form chitin microfibrils in vitro, and it has been suggested that in vivo they transport chitin synthase to the plasma membrane at the hyphal apex and at the positions of developing septa, which are the places where chitin synthesis is highly localised in actively growing vegetative hyphae (Bartnicki-Garcia, 2006).

Using chitin synthases from Neurospora crassa labelled with green fluorescent protein, and high-resolution confocal laser scanning microscopy of living hyphae, Riquelme et al. (2007) and Riquelme & Bartnicki-García (2008) studied the trafficking of chitin synthase during hyphal growth. They showed that at the hyphal apex, fluorescence label was localised to a single conspicuous body (the Spitzenkörper), whilst at distances more than 40 µm from the apex the chitin synthase fluorescence occurred in a network of endomembrane compartments, mostly irregularly tubular, but some spherical. In between (20 to 40 µm from the apex), fluorescence occurred as globules that disintegrated into vesicles as they moved forward and ultimately contributed to the Spitzenkörper. These in vivo observations suggested that Spitzenkörper fluorescence specifically:

' …originated from the advancing population of microvesicles (chitosomes) in the subapex.' (Riquelme et al., 2007).

It was also demonstrated that brefeldin A (known to be a specific inhibitor of protein transport in Golgi dictyosomes) was unable to interfere with the traffic of chitosomes, and that fluorescence associated with the labelled chitin synthase was not associated with the endoplasmic reticulum. These observations support the idea that chitin synthase proteins are delivered to the cell surface by a secretory pathway distinct from the classic endoplasmic reticulum-to-Golgi. The finding of a direct interaction between chitin synthase proteins equipped with their own myosin motors and the actin cytoskeleton (Takeshita et al., 2007) also supports this interpretation.

Once delivered to its site of action, chitin synthase is an integral membrane-bound enzyme (a transmembrane protein). The enzyme protein spans the membrane, accepting substrate monomers from the cytoplasm on the inner face of the membrane and extruding the lengthening chains of chitin through the plasma membrane as they are made. Hydrogen bonding between the newly formed polymers of chitin results in microfibril formation and subsequent crystallisation of chitin in the extracellular space immediately adjacent to the plasma membrane. This process of chitin synthesis primarily occurs at sites of active growth and cell wall remodelling (Gooday, 1995b). Although during normal apical growth of the hypha the incorporation of newly synthesised chitin is limited to the hyphal apex, there is evidence that inactivated chitin synthases are widely distributed in the plasma membrane. One piece of evidence for this is that inhibition of protein synthesis in Aspergillus nidulans resulted in chitin synthesis occurring uniformly over the hypha. This observation implies that protein synthesis is required to maintain the inactivity of chitin synthases already in place in the membrane. Inactivated chitin synthase are an intrinsic component of the mature hyphal plasma membrane, the enzyme(s) being activated somehow specifically at the hyphal apex and at sites where branch or septum formation is initiated.

Chitin is important at particular sites in yeast walls, although the major structural component in these organisms is a fibrillar inner layer of β-glucan. In Saccharomyces cerevisiae, chitin is concentrated mostly in septa. This yeast has three chitin synthases (Chs1, Chs2, and Chs3). Chs3 is a class IV chitin synthase, and is required for formation of chitin rings at the base of emerging buds and for chitin synthesis in the lateral cell wall during vegetative growth. Chs3 is found at the plasma membrane and in chitosomes, and its correct localisation requires the functioning of several other proteins. The Chs3 chitin synthase is responsible for generating approximately 80–90% of the total chitin of the yeast cell. Chs2, a class II enzyme, synthesises chitin in primary septa. Appearance of Chs2 is cell cycle-dependent; it appears at a late stage of mitosis and is localised to the septation site and degraded immediately after septum formation. Chs1, a class I enzyme, repairs the weakened cell walls of daughter cells after their separation from mother cells; this separation is executed by the activity of a chitinase that digests the primary septa; Chs1 chitin synthase repairs the wall, replenishing chitin polymers lost during cytokinesis. Chs1 exists at a constant level throughout the cell cycle and is present in the plasma membrane and chitosomes. The simultaneous deletion of all genes coding for these three chitin synthases is lethal for the cell, demonstrating that chitin is an indispensable component of the cell wall of S. cerevisiae even though it is a minor component (Bowman & Free, 2006; Lesage & Bussey, 2006).

Updated August, 2019