Regulation of expression, activity and localization of fungal chitin synthases

Abstract

The fungal cell wall represents an attractive target for pharmacologic inhibition, as many of the components are fungal-specific. Though targeted inhibition of β-glucan synthesis is effective treatment for certain fungal infections, the ability of the cell wall to dynamically compensate via the cell wall integrity pathway may limit overall efficacy. To date, chitin synthesis inhibitors have not been successfully deployed in the clinical setting. Fungal chitin synthesis is a complex and highly regulated process. Regulation of chitin synthesis occurs on multiple levels, thus targeting of these regulatory pathways may represent an exciting alternative approach. A variety of signaling pathways have been implicated in chitin synthase regulation, at both transcriptional and post-transcriptional levels. Recent research suggests that localization of chitin synthases likely represents a major regulatory mechanism. However, much of the regulatory machinery is not necessarily shared among different chitin synthases. Thus, an in depth understanding of the precise roles of each protein in cell wall maintenance and repair will be essential to identifying the most likely therapeutic targets.

The fungal cell wall is a complex cross-linked network of chitin, glucans, other polysaccharides as well as integral proteins. The central core consists of glucans cross-linked to chitin, with various decorating polysaccharides depending on the species (reviewed in [1]). Many of the components of the cell wall are fungal-specific, thus inhibition of these components represents a logical target for antifungal agents. This approach has been validated by the echinocandin class of antifungals that target β-1,3 glucan synthesis [2–6]. However, the fungal cell wall is a dynamic and developmentally plastic construction, capable of compensating for loss of β-1,3-glucan by increased chitin deposition [7–11].

Chitin is a linear homopolymer of β-1,4-linked N-acetylglucosamine (Glc-NAc). Chitin fibrils occur in different conformations, both as long thin microfibrils and as short thick rodlets, suggesting that specific forms of chitin may be important in different structural roles [12]. Chitin can also be deacetylated to chitosan, a more flexible and soluble polymer, important in Saccharomyces cerevisiae ascospores and in the cell wall of Cryptococcus neoformans [13, 14]. In addition, the pattern of cross-linking to other polysaccharides varies depending on the type of chitin microfibril as well as its subcellular location, consistent with a highly organized and regulated cell wall structure [12, 15]. Chitin biosynthesis is limited to fungi and insects, thus chitin biosynthesis represents an as yet unexploited target for therapeutic intervention in fungal disease affecting humans.A number of chitin synthesis inhibitors have been identified, the best characterized of which are the nikkomycins. Nikkomycins are natural products of Streptomyces tendae that are competitive inhibitors of chitin synthase enzymes (reviewed in [16]). Enzyme kinetic studies in the genetically tractable yeasts have showed wide discrepancy in activity of the inhibitor against the various enzymes. In both S. cerevisiae and Candida albicans, nikkomycin Z is much less active against the chitin synthases that fill near-essential roles in yeast cell wall biosynthesis [17–19]. Studies of nikkomycin Z treatment of murine models of candidiasis, histoplasmosis, blastomycosis, and aspergillosis have discordant results. Nikkomycin Z monotherapy of dimorphic infections showed improved survival and some microbiologic cures [20–22]. In contrast, mice with candidiasis did have better survival while on treatment, but all relapsed off of therapy, suggesting that nikkomycin Z may not be fungicidal in Candida [23]. In a model of invasive aspergillosis, nikkomycin Z was not effective as monotherapy [24]. However, it did appear to show some synergy with other agents, particularly echinocandins, raising the possibility of improved treatment with simultaneous targeting of two cell wall components [24–27]. This result is supported by in vitro data that demonstrates markedly enhanced cell wall damage to fungal cultures grown in the presence of both chitin and glucan inhibitors as compared to cultures grown with one or the other [28, 29]. Chitin synthase inhibitors have not been employed clinically, though there has been a recent resurgence of interest in these agents, particularly for dimorphic fungal infections. In fact, the first human phase I pharmacokinetic study of nikkomycin Z was recently published and there are plans for a phase II efficacy trial of treatment of coccidioidal pneumonia [30, 31].

In this review, we have focused on the regulation of the chitin synthases, particularly in model species or medically relevant fungal pathogens, though significant contributions to our understanding of chitin synthesis regulation have also been drawn from research of agricultural fungal pathogens. The role of chitin in cell wall structure and pathogenicity have been recently reviewed [1, 32].

The role of individual chitin synthases cannot be predicted based on homology

Chitin is an important structural component in the septa and cell walls of fungi, but depending on the fungal species, chitin can compose from a small minority up to nearly half of the cell wall dry weight [33]. In yeast, chitin is a minor cell wall component, totaling only 1–2% of the dry weight [34] and primarily found as a minor component in the lateral walls as well as concentrated in the bud neck, bud scar and septa. The situation in filamentous fungi is different, as chitin tends to be a much larger overall cell wall component (up to 40% or more of the dry weight) [35, 36] and is more diffusely distributed in the cell wall with increased deposition at the hyphal tips and septa. In S. cerevisiae, in which chitin is a minor cell wall component, strains containing simultaneous disruption of all three chitin synthases are nonviable except in the setting of acquisition of suppressor mutations, affirming the essential nature of chitin in fungi [37, 38].

The chitin synthases are integral plasma membrane proteins that catalyze polymerization of UDP-Glc-NAc into hydrophobic chitin chains that are then extruded through the cell membrane and incorporated into the cell wall [39, 40]. Chitin synthase (CHS) enzymes are encoded by members of a large gene family (Figure 1A), suggesting the possibility of both functional specialization as well as redundancy. Different chitin synthases produce chitin that is localized to specific cell wall derived structures or developmental stages [12]. Phylogenetic analysis demonstrates the existence of seven classes of chitin synthases divided into two families (Figure 1A). Class I, II and IV genes are present in all fungi, whereas classes III, V, VI and VII are specific to filamentous fungi and certain dimorphic species (Figure 1B). The number of putative chitin synthase genes within each species varies, with three in S. cerevisiae, four in C. albicans, seven in Wangiella dermatitidis and Neurospora crassa and eight each in Aspergillus nidulans, A. fumigatus and C. neoformans (Table 1).

Figure 1. Chitin synthase gene functional domains.

A. Phylogenetic analysis of the chitin synthases of A. fumigatus, A. nidulans, C. albicans, C. neoformans, N. crassa, S. cerevisiae and W. dermatitidis demonstrates seven classes of chitin synthases, divided into two families. Yeasts contain enzymes from classes I, II and IV, whereas filamentous fungi tend to contain at least one enzyme from each class.

B. The proteins encoded for by the class I, II and III genes tend to be smaller, and contain an amino terminal catalytic domain and multiple carboxy transmembrane domains. The proteins encoded by class IV, V, VI and VII genes tend to be larger and the catalytic domain tends to be more towards the carboxy terminal. The proteins encoded by class V and VII genes usually contain an amino terminal domain with some homology to myosin motor domains and a carboxy terminal with chitin synthase catalytic residues.

Table 1.

Characteristics of chitin synthase genes among model and pathogenic yeast and molds

Spewcies	Gene	Class	Mutant phenotype	Transcriptional changes	Protein localization	References
Af	chsA	I	No phenotype	Increased with echinocandin	ND	[48, 89, 141]
Af	chsB	II	No phenotype	ND	ND	[48, 141]
Af	chsC	III	No phenotype	Increased with echinocandin	ND	[48, 89, 141]
Af	chsD	VI	Decreased chitin	ND	ND	[77, 141]
Af	chsE	V	Decreased chitin, hyphal defects, normal virulence	ND	ND	[75, 141, 142]
Af	chsF	IV	No phenotype	ND	ND	[141]
Af	chsG	III	Hyphal defects, poor growth and conidiation, less virulent	ND	ND	[77, 142]
Af	chsEb	VII	ND	ND	ND	[143]
An	chsA	II	Decreased conidiation	Asexual development	Septation sites	[68, 73, 132, 133, 144– 149]
An	chsB	III	Severe growth defect	Constitutively expressed	Germ tubes, hyphal tips, septa, polarized growth sites, asexual structures	[56–58, 73, 145, 147]
An	chsC	I	None	Hyphae and conidiophores	Hyphal tips, septation sites	[47, 57, 68, 132, 145– 149]
An	chsD	IV	Normal chitin or decreased chitin	Hyphae and conidiophores	ND	[57, 68, 69, 74, 148– 150]
An	chsF	III	ND	ND	ND	[151]
An	chsG	VI	ND	ND	ND	[151]
An	csmA	V	Poor conidiation, Abnormal hyphae	Changes in osmolarity	Hyphal tips, septation sites, binds actin	[69, 74, 78, 136, 149, 152, 153]
An	csmB	VII	Impaired hyphal growth, poor conidiation	ND	Hyphal tips, forming septa, binds actin	[78, 138]
Ca	CHS1	II	Lethal	chs deletions, cell wall stress	Mother-bud neck, vacuoles	[12, 45, 52, 81, 83, 154]
Ca	CHS2	I	Slower growth, decreased chitin	Cell wall stress, hyphal induction	Mother-bud neck, Peripheral patches, vacuoles	[12, 42, 45, 65, 81, 83]
Ca	CHS3	IV	Decreased rodlet chitin, decreased virulence	chs deletions, cell wall stressors, hyphal induction	Mother-bud neck, septa, small cytoplasmic patches	[12, 45, 64, 65, 83]
Ca	CHS8	I	No long chain microfibrils	Certain otherchs deletions, cell wall stress	Bud-neck, septa, Cytoplasmic patches, hyphal tip	[12, 46, 83]
Cn	CHS1	IV	None	Lower	ND	[49]
Cn	CHS2	III	None	Highest	ND	[49]
Cn	CHS3	IV	Temperature lethal, growth defects, stress sensitivity, decreased chitin	Highest	ND	[49]
Cn	CHS4	VII	None	Lower	ND	[49]
Cn	CHS5	V	Temperature sensitivity	Highest, certain other chs deletions	ND	[49]
Cn	CHS6	I,II	Temperature sensitivity	Very low	ND	[49]
Cn	CHS7	III	Temperature sensitivity	Absent except with chs3 deletion	ND	[49]
Cn	CHS8	I,II	Temperature sensitivity	Highest	ND	[49]
Nc	chs-1	III	Abnormal hyphae	ND	Hyphal apex, developing septa	[61, 98]
Nc	chs-2	II	None	ND	ND	[53]
Nc	chs-3	I	ND	ND	Hyphal apex,septa	[97]
Nc	chs-4	IV	None	ND	ND	[70]
Nc	chs-5	V	ND	ND	ND	[129]
Nc	chs-6	VI	ND	ND	Hyphal apex,septa	[97]
Nc	chs-7	VII	ND	ND	ND	[129]
Sc	CHS1	I	Bud lysis defect	Increased during sexual cycle	ND	[17, 37, 41, 43, 44, 54, 90]
Sc	CHS2	II	Aseptate	Decreased during sexual cycle	Bud neck during ring contraction	[37, 51, 54, 101, 102, 117]
Sc	CHS3	IV	Decreased cell wall chitin, defective spore maturation	Vegetative growth and septum formation	Endocytic vesicles, bud site, small and large bud neck, plasma membrane with stress	[14, 62, 63, 101, 104, 106]
Wd	CHS1	II	Failure to separate, broader mother- bud neck	Other chs deletions	ND	[55, 84]
Wd	CHS2	I	No phenotype	chs deletions	ND	[50, 55, 84]
Wd	CHS3	III	No phenotype	chs deletions	ND	[50, 60, 84, 155]
Wd	CHS4	IV	Reduced chitin, abnormal yeast clumping	Constitutive, increases with stress,chs deletions	ND	[71, 84]
Wd	CHS5	V	Suppressive growth defects, less virulent	Cell wall stress	Yeast buds, hyphal tips, sites of expansion, actin-dependent	[76, 84, 137, 156]
Wd	CHS6	VI	ND	ND	ND	[137]
Wd	CHS7	VII	ND	ND	ND	[137]

Af: Aspergillus fumigatus; An: Aspergillus nidulans; Ca: Candida albicans; Cn: Cryptococcus neoformans; Nc: Neurospora crassa; Sc: Saccharomyces cerevisiae; Wd: Wangiella dermatitidis; ND: not done.

Homology-based prediction of specific chitin synthase function is highly imperfect, as individual classes do not necessarily have the same function in different species. To date, most research has employed classical genetic techniques, primarily examining the phenotype of single and multiple gene disruptions. Extension of mutant phenotypes into functional descriptions is complicated by the existence of multiple enzymes with possible functional redundancy. In addition, extrapolation from yeast to filamentous fungi is necessarily limited by the widely differing nature of their cell walls and life cycles.

Class I CHS genes were the first to be described, as they are highly active in chitin synthase activity assays in vitro, though their actual contribution to chitin synthesis is relatively small [41, 42]. Functionally, S. cerevisiae Chs1p acts as a repair enzyme at the site of cytokinesis, counterbalancing the activity of chitinase in mediating separation, thereby maintaining cell wall integrity [43, 44]. There are two class I chitin synthase genes in C. albicans, CHS2 and CHS8. Both Chs2p and Chs8p appear to have modest effects on overall growth and cell wall chitin content, and unlike S. cerevisiae, no bud lysis defect has been elicited even when both genes are disrupted [42, 45, 46]. Mutation of the class I genes in A. nidulans, A. fumigatus and C. neoformans does not yield any obvious phenotypes, although mutation of CHS2 of W. dermatitidis results in decreased chitin synthase activity [47–50].

Class II genes also tend to make a relatively small contribution to total cellular chitin, but functionally are quite important in S. cerevisiae and C. albicans, but not A. nidulans or N. crassa [47, 51–53]. S. cerevisiae Chs2p acts in construction of the primary septum after cytokinesis [37, 54]. C. albicans Chs1p is essential, owing to its role in producing both septal and lateral wall chitin [52]. The W. dermatitidis class II chs1Δ strain is unable to transition between growth types and yeast cells fail to divide normally, though rather than absent septa as seen in the yeast mutants, there is a broader neck with enriched chitin content at the septa [55].

Class III genes appear to be relatively important for normal growth and morphology in Aspergilli. Both A. nidulans and A. fumigatus each contain two class III genes. A. nidulans ChsB appears to play a role in synthesis of chitin in the hyphal tips and conidia, owing to marked defects in hyphal morphology and poor conidiation when mutated [56–58]. Deletion of A. fumigatus chsG alone or both class III genes (chsC and chsG) produces similar but less marked defects, suggesting that the functions may be related to those proposed in A. nidulans [59]. Disruption of the homologous genes in W. dermatitidis yields decreased chitin synthase activity but no apparent growth defects and the C. neoformans mutant has no phenotypes [49, 60]. Mutation of N. crassa chs-1 alters hyphal morphology and decreases overall activity [61].

Class IV chitin synthases are the bulk chitin synthases in yeasts. The C. albicans and S. cerevisiae class IV genes encode proteins that produce the majority of the total cell wall chitin, but loss of these genes does not produce equally dramatic phenotypes. S. cerevisiae CHS3 (also called CAL1, CSD2 and DIT101) encodes the chitin synthase responsible for the majority of overall chitin content [14, 62, 63], similar to the role of C. albicans CHS3 [45, 64–66]. Yeast class IV genes appear to function in the cell wall compensatory response to cell wall stressors, as double mutants with S. cerevisiae GAS1, a β-1,3-glucanosyltransferase that modifies cell wall cross-linking, and the Chs3p regulators CHS4, CHS5, CHS6 and CHS7 result in osmotic support-dependent viability [67].

However, the generalization of the class IV chitin synthases as bulk synthases cannot be extended to other fungi as the mutant phenotypes of the corresponding genes in filamentous fungi are not similar to the yeasts or even to each other. In A. nidulans, loss of chsD results in no defects in cell growth or morphology and no decrease or a modest decrease in chitin content [68, 69] and in N. crassa, the chs-4^RIP strain does not have any phenotypes [70]. Slightly more similar to the yeasts, the W. dermatitidis chs4Δ strain contains decreased chitin [71]. In contrast, the C. neoformans chs3Δ strain is slow-growing and more sensitive to cell wall damage as well as producing enlarged abnormally shaped cells with increased chitin and reduced chitosan content [49]. The ability of this mutant strain cell wall to retain melanin was reduced, suggesting that melanin may associate with chitin microfibrils synthesized by specific chitin synthases [49], which has also been seen in C. albicans [72].

Disruption of class V genes in A. nidulans, A. fumigatus and W. dermatitidis produces fairly dramatic changes in growth and morphology, but the same is not seen in C. neoformans [49, 69, 73–76]. For instance, A. nidulans CsmA functions in hyphal growth as well as septal formation as evidenced by the mutant phenotypes, and strains deficient in A. fumigatus chsE have similar but less severe defects in hyphal growth [69, 73–75]. The W. dermatitidis chs5Δ strain has defective growth and morphology and even ceases to be viable when grown without osmotic support at elevated temperatures, with an accompanying decrease in virulence [76].

Among class VI genes, disruption of A. fumigatus chsD results in moderately decreased total chitin content without change in chitin synthase activity or accompanying defects in growth or morphology [77].

Loss of the class VII gene A. nidulans csmB results in the formation of ballooned and intrahyphal hyphae and increased tendency towards of lysis of subapical regions [78]. A. nidulans ΔcsmA ΔcsmB is synthetically lethal suggesting that the myosin motor domain-containing chitin synthases play an important role in chitin synthesis in filamentous fungi species [78].

It is clear from the chitin synthase gene disruption studies that prediction of the function of individual genes is not reliable based on homology to other genes of the same class from other fungal species. Importance of chitin synthases may thus lie with their highly specialized roles in very specific aspects of fungal growth and development.

Cell cycle and developmental stage alter yeast chitin synthase transcription

In yeast, chitin synthase gene transcription changes throughout the cell cycle as well as during different developmental stages. Transcription of S. cerevisiae CHS1 peaks in M/G1, CHS2 in M-phase and CHS3 after septum formation and before division, which is consistent with the role of Chs1p in repair of the bud scar after cytokinesis and Chs2 in primary septum formation [14, 79]. In C. albicans, transcription of CHS1, CHS8 and CHS3 rise in G2, whereas CHS2 does not demonstrate cell cycle dependent changes [80]. In terms of developmental stage differences in transcription, C. albicans CHS1 is expressed at low levels in early germ tube development and equally in yeast and hyphal forms [45, 81]. C. albicans CHS2 transcription increases during hyphal growth [45, 81], consistent with its role in septum formation. C. albicans CHS3 expression increases in the transition from yeast to hyphal growth, and then diminishes thereafter [45, 82]. This linkage of individual chitin synthase transcription to specific developmental stages argues that chitin biosynthesis may not just serve as general cell wall structural support, but may also be closely regulated in ways that allow specific remodeling of the cell wall as needed throughout growth. Similar studies have not been done in filamentous fungi, as the vastly different development program of filamentous fungi renders cell wall development less tied to the cell cycle.

Transcriptional compensation occurs with disruption of chitin synthase genes, suggesting functional redundancy between different chitin synthase proteins

The large number of chitin synthase genes, particularly in filamentous fungi, raises the question of why so many genes are present: it may reflect functional redundancy. In both yeast and filamentous fungi, compensatory increases in transcription have been found in response to disruption of chitin synthase genes. Transcription of C. albicans CHS1 increases with disruption of nearly every other chitin synthase, consistent with its functional importance, and CHS2 is not upregulated in any of the mutants [83]. In W. dermatitidis, all other single gene deletions result in increased CHS5 transcription, though chs5Δ does not cause any changes in transcription of the other chitin synthase genes [84]. Thus, compensatory transcriptional increases of chitin synthase genes occurs in fungal strains carrying one chitin synthase gene disruptions, but the pattern of upregulation is mutation-dependent.

Disruption of the cell wall results in increased chitin synthase gene transcription, mediated via several different signaling pathways

The cell wall integrity (CWI) pathways encompass a number of compensatory cell wall changes that occur in response to treatment with cell wall perturbing agents or mutations in cell wall synthetic or regulatory genes. Mutations in various cell wall genes result in global changes in S. cerevisiae gene transcription, particularly among components of a variety of cell signaling pathways, including the protein kinase C mitogen-activated protein (PKC-MAP) kinase pathway, the Ca²⁺/calcineurin pathway and a generalized global stress response pathway [11]. It is important to note that the specific response, including the pattern of alterations in gene transcription, varies depending on the specific stressor [85, 86]. In C. albicans, the high osmolarity glycerol (HOG) response MAP kinase pathway and the Ca²⁺/calcineurin pathway have also been implicated in the compensatory reaction to cell wall stressors [83]. C. albicans CHS gene expression analysis in the setting of various inducers of the CWI pathway results in upregulation of different combinations of chitin synthases depending on the treatment [83]. Mutations in components of the Ca²⁺/calcineurin pathway results in increased transcription of chitin synthases, though the pattern differs depending on which component is deficient, whereas mutations in components of the PKC and HOG pathways cause generally decreased CHS transcription [83].

C. albicans CHS1 transcription is regulated by PKC, HOG and Ca²⁺/calcineurin signaling. Specific promoter elements mediate transcriptional control by the PKC pathway, but deletion of consensus motifs for Ca²⁺/calcineurin and HOG pathway transcriptional control does not correlate with the loss of expression [87]. Thus, the basis for transcriptional control of CHS1 by these pathways may not necessarily be through the classically defined downstream transcription factors. In addition, transcriptional responses for other chitin synthases have been studied, but whether the identified responses are mediated via stereotypical transcription factors remains undefined at this time.

Treatment of C. albicans with echinocandins, noncompetitive inhibitors of β-1,3-glucan synthesis, results in increased transcription of all four chitin synthase genes [88]. Conversely, ectopic activation of these pathways permits survival of C. albicans in the presence of otherwise lethal doses of echinocandins and suggests that these pathways could provide a target for synergistic targeting of multiple cell wall components [88]. Mutations in the HOG, PKC and Ca²⁺/calcineurin pathways prevent this upregulation, though disruption of each pathway produces a different pattern of transcriptional increases [88]. These pathways may each regulate subtly different aspects of cell wall compensatory responses. Similar echinocandin-mediated transcriptional upregulation is seen for two chitin synthases in A. fumigatus, which is abrogated by calcineurin inhibition or mutation [89], suggesting that inhibition of these compensatory pathways may represent a possible future antifungal targeting strategy.

Post-transcriptional regulation of chitin biosynthesis is important for modulation of chitin synthase activity

Changes in chitin synthase gene transcription do not necessarily correlate with changes in cellular chitin content or chitin synthase activity, thus regulation of chitin synthesis likely occurs both transcriptionally and post-transcriptionally. In S. cerevisiae, blocking CHS2 transcription results in quicker loss of Chs2p activity than loss of CHS2 mRNA, suggesting that the Chs2p protein is short-lived [90]. S. cerevisiae CHS1 has cyclical mRNA transcription but Chs1p activity is constant throughout the cell cycle [90]. S. cerevisiae Chs3p has prolonged activity after cessation of gene transcription suggesting that protein is stable and long-lived [90].

Studies utilizing chitin synthase activity assays must be cautiously interpreted, given the likelihood that in vitro conditions of the assay may not mirror in vivo conditions, possibly explaining why significant discordance has been found between total chitin synthase activity and total cellular chitin content. In addition, the existence of large chitin synthase gene families makes it difficult to identify the quantity and quality of chitin ascribable to individual chitin synthases. Historically, chitin synthases have been grouped as zymogenic or non-zymogenic based on increased chitin synthase activity in cell membrane extracts after addition of proteases thus regulation may occur by proteolytic cleavage [91]. To date, no convincing candidate protease has been identified and it is unclear if proteolysis is a true regulatory mechanism in vivo. Expression of a truncated S. cerevisiae Chs2p result in higher protein levels in vivo, but in vitro activity from both the full-length and truncated proteins are similar [92]. Heterologously expressed S. cerevisiae Chs2p can be hyperactivated by an extract containing an undefined soluble yeast protease activity but this may not mirror actual in vivo conditions [93].

Regulation of chitin synthase activity and overall chitin deposition is affected by cell wall stress

A number of different disruptors of cellular homeostasis, both directly active on the cell wall and others that are more generalized, have been shown to alter chitin synthase activity and cell wall chitin content, synthesized by different chitin synthases depending on the trigger. Mutation of the S. cerevisiae β-1,3-glucan synthase results in increased, though delocalized, cell wall chitin associated with both increased Chs1p and Chs3p activity [7]. Pharmacologic inhibition of glucan synthesis in C. albicans also produces increased chitin synthase activity and cell wall chitin content [88], demonstrating the dynamic ability of fungi to respond to cell wall damage. A number of different cell membrane associated proteins have been implicated in these responses. S. cerevisiae Mid2p is a plasma membrane associated cell wall sensor that mediates cell wall integrity via induction of Pkc1p MAP kinase signaling. Mutation of MID2 leads to decreased chitin deposition in response to cell wall stress, a response that is dependent on the presence of intact Chs3p [94]. Loss of GGP1/GAS1, which encodes a glycosylphosphatidylinositol-anchored plasma membrane glycoprotein, decreases cell wall glucan and increases chitin, dependent on Chs3p, suggesting a central role for CHS3 in mediating cell wall stress compensatory responses [8, 10]. In contrast, strains missing Bgl2p, a cell wall protein of unclear function; increase Chs1p activity as well as cell wall chitin content in a CHS3-dependent manner and Chs1p activity [95]. Whereas many of the studies have suggested a central role for Chs3p in this compensatory response, other chitin synthases also may function similarly raising the question of whether various cell wall disruptions acts through different signaling pathways to the same end result.

The same signaling pathways that have been shown to regulate chitin synthase transcription have been shown to lead to alterations in chitin synthase activity and total cell wall chitin content. However, the net effects on transcription do not necessarily align with the changes in either activity or content, highlighting both the limitations of the assays and the likelihood that regulation of this process occurs at multiple stages. In general, external cell wall stressors and blocking of signaling via pathways implicated in the response to these stressors result in increased chitin synthase activity and/or increased chitin content. Calcium treatment of C. albicans leads to increased Chs2p and Chs8p activity and increased Chs3p-dependent chitin content [83, 88], again highlighting the lack of correlation between measurable activity and total content. This stress response is important for survival, as calcium and calcofluor white pretreatment of the normally nonviable C. albicans chs1Δ chs3Δ strain facilitates formation of a salvage septum [88]. This reinforces the inherent complexity of antifungal agent design and suggests the possibility that simultaneous inhibition of multiple targets may represent a more clinically effective mechanism.

Plasma membrane localization of chitin synthases occurs via specialized secretory pathways

Plasma membrane extracts contain intrinsic chitin synthase activity [39]. Membrane fractionation yields separate vesicular populations containing chitin synthase activity, one associated with intracellular specialized secretory vesicles, deemed chitosomes, and the other associated with the plasma membrane [40]. The presence of intracellular vesicles is consistent with the existence of reserves of chitin synthesis that may be accessible as a functional reservoir via regulated localization to the plasma membrane. However, the processes that mediate the recruitment of chitin synthases to the plasma membrane are as yet undefined.

Extension of fungal hyphae depends on the action of cell wall biosynthetic enzymes, including chitin synthases, at the hyphal apex [96]. Chitin synthase localization to the plasma membrane has been shown to occur via a highly specialized secretory apparatus, distinct from the classic secretory pathway [97]. Transport appears to depend on certain cytoskeletal structures, especially the actomyosin skeleton [98]. Different chitin synthases are transported via different pools of vesicles, and these vesicles do not transport other cell wall biosynthetic enzymes [40, 98]. Vesicles fuse with the membrane at sites of active cell wall growth such as septa and hyphal tips. Based on data from N. crassa, this process is thought to occur in a two-step process, in which the enzyme moves through its specialized secretory apparatus to a collection of vesicles near the hyphal apex that may function as a reserve for supplying growth [99, 100]. From there, the vesicles move to the plasma membrane. For yeast chitin synthases, some may be recycled back into the vesicles whereas others are synthesized, localized and degraded in a cell cycle dependent manner [101, 102].

N. crassa chitin synthases cluster in vesicles near the hyphal apex, as well as occurring at lower levels in the hyphal tip plasma membrane [103]. Tagged N. crassa class I and VI chitin synthases have been traced from distal endomembranous compartments proximally to smaller globular bodies and finally to vesicles that concentrate at the Spitzenkörper [97]. Tagged class III chitin synthases partially co-localizes with the other tagged chitin synthases, but are found in different vesicles, consistent with specific secretory processes for individual chitin synthases [98]. In addition, this localization was dependent on actin but not microtubules [98], implicating cytoskeletal interactions in this process.

The yeast chitin synthase 3 protein secretory mechanism depends on multiple proteins

Detailed molecular biologic analysis of chitin synthase localization has been studied only in yeast. As a consequence, the secretion of some chitin synthases is fairly well understood, but it is not clear to what extent this can be generalized, given the larger number of chitin synthase genes and the presence of classes specific to filamentous fungi.

Studies of S. cerevisiae Chs3p suggest that sequestration of chitin synthases to internal compartments and thereafter to the plasma membrane is a highly regulated process (Figure 2). Chs3p is normally polarized to a diffuse ring at the future bud site and the neck of small buds in a myosin-dependent manner [104]. A proportion of Chs3p is contained in internal compartments that are endocytically derived, suggesting that Chs3p may be shuttled into and out of the membrane to titrate its activity [101]. The Rab protein Ypt32p, a mediator of vesicle fusion with the plasma membrane, regulates Chs3p delivery to the plasma membrane [105]. Various cell wall stressors trigger the redistribution of Chs3p from chitosomes to the plasma membrane, suggesting a mechanism through which fungi may rapidly adjust chitin synthesis in response to stress [106]. The re-localization of Chs3p can be blocked by mutations in the regulatory GTPase Rho1p and the MAP kinase Pkc1p [106]. In addition, Chs3p has been shown to be phosphorylated by Pkc1p and redistribution of Chs3p from internal stores to the plasma membrane occurs with expression of constitutively active Pkc1p [106]. In C. albicans, Chs3p is found at the mother-bud neck and in small cytoplasmic patches during yeast growth, as well as at the septa and tips during hyphal growth [12]. This localization depends on the ability to alter the phosphorylation status of serine 139 [107]. However, loss of PKC1 does not affect Chs3p localization in C. albicans, in contrast to the role played in S. cerevisiae [106, 107].

Figure 2. Transit of Chs3p from the ER to the plasma membrane.

The movement of S. cerevisiae Chs3p from the ER to the plasma membrane requires both direct and indirect interactions with a number of specialized proteins. The thick arrow represents the path followed by Chs3p. Thin arrows indicate proteins whose functions promote Chs3p transit. Chs7p is an ER protein that may facilitate Chs3p folding [108]. Pfa4p palmitoylates Chs3p [110]. Chs5p facilitates assembly of a multi-protein complex containing Chs6p that binds Chs3p to allow movement through the trans-Golgi [104, 111, 112, 114, 115]. Mutations in the signaling pathway genes PKC1, RHO1, and PIK1, the vesicle fusion mediator YPT32, and cytoskeletal gene MYO2 prevent plasma membrane localization of Chs3p [104–106]. Generation of phosphoinositide phosphate via the Pik1p kinase is important for transport from the ER to the plasma membrane, with negative regulation of this process mediated via the Sac1p phosphatase [116].

Other proteins that play a role in mediating Chs3p localization have been identified as a consequence of the marked drop in chitin content in these mutant strains. These proteins are not chitin synthases per se, but instead interact with Chs3p to regulate its trafficking through the secretory machinery and eventual localization to the plasma membrane. S. cerevisiae Chs7p is an endoplasmic reticulum (ER) membrane protein whose mutations results in ER retention of Chs3p [108]. Deletion of C. albicans CHS7 also leads to diminished chitin content and abnormal septation [109]. Expression of S. cerevisiae CHS7 is limiting for Chs3p activity, and many of the developmental processes and external factors that increase cell wall chitin result in increased CHS7 transcription [108]. Pfa4p, a protein palmitoyltransferase, is also required for Chs3p to exit the ER, though Pfa4p and Chs7p appear to function independently [110].

Other proteins have been identified that are important for transport of the class IV chitin synthases through later secretory stages. S. cerevisiae Chs5p and Chs6p are important for Chs3p passage through the trans-Golgi [104, 111, 112]. S. cerevisiae Chs1p is shuttled via the endocytosis pathway and its localization is not affected by loss of CHS6, confirming that regulation is not generalizable between different chitin synthases [111, 113]. Chs5p colocalizes with Chs3p in cytoplasmic patches, and loss of CHS5 results in retention of Chs3p in these patches and absence of subsequent plasma membrane localization [104]. Chs6p belongs to a four member gene family in which the separate member proteins bind each other and Chs5p, after which they associate with Chs3p to facilitate export from the Golgi [114, 115]. This specialized secretory process is dependent on the actomyosin skeleton, as disruption of myosin results in delocalized Chs3p [104]. As with many other steps in the localization of chitin synthesis, phospho-signaling plays an important regulatory role. Loss of Sac1p, a PI phosphatase and overexpression of Pik1p, a PI-4-kinase, results in abnormal accumulation of Chs3p in vacuoles, thus this phosphorylation pathway may function as a regulator of transport from the Golgi [116].

Chs3p interacts with an accessory protein as well as other scaffold proteins to maintain activity and plasma membrane localization

Rather than regulating transit of Chs3p through its specialized secretory machinery, Chs4p is a Chs3p activator. Loss of CHS4 (also known as CAL2, SKT5 and CSD4) in S. cerevisiae results in decreased cell wall chitin due to diminished Chs3p activity [62, 66, 117–119], but does not mediate its activity via CHS3 transcription or translation [120]. Instead, it appears that a direct physical interaction between farnesylated Chs4p and Chs3p is important for its function [121–123]. Chs3p can localize to the bud neck without Chs4p, but is inactive and is rapidly endocytosed; thus Chs4p may function to maintain Chs3p at the correct plasma membrane site in a stable and activated form [122].

Chitin synthase 3 is one of the components of a protein complex at the mother bud neck that positions the primary septum (Figure 3). Much of the work towards defining this complex in S. cerevisiae has focused on Bni4p, a scaffold protein that facilitates the interaction of septal filaments to an assortment of proteins at this complex, including the Chs3p activator Chs4p. The chitin synthase is likely recruited to the bud neck after the eventual septum position has been determined, given the septin- and Bni4p-dependent localization of Chs3p and Chs4p [124]. Bni4p directly interacts with the type 1 serine-threonine phosphatase Glc7p; if this does not occur, then Bni4p does not bind Chs4p and Chs3p is not localized to the bud neck [125, 126]. The targets for Glc7p activity at the bud neck complex are not known; there is no evidence for activity towards Chs3p, Chs4p or the septins, perhaps Bni4p itself may be the target. Bni4p is phosphorylated in a cell cycle-dependent fashion with loss of the Hsl1p kinase resulting in increased Bni4p at the bud neck; Hsl1p may decrease the Bni4p-Glc7p interaction [125]. Assembly of the chitin ring and primary septum assembly is a complicated process that requires the coordinated interaction of a variety of proteins, and future additional mechanistic understanding may permit identification of possible novel therapeutic targets.

Figure 3. Localization of Chs3p to the bud neck.

Localization of Chs3p to the mother-bud neck requires a protein complex containing septins, Bni4p, Glc7p and Chs4p [124–126]. Chs3p can reach the bud neck without Chs4p [121–123], but activity and sustained localization of Chs3p depends on all of the proteins. The target of the Glc7p phosphatase is not known, and the kinase Hsl1p may negatively regulate the interaction between Glc7p and Bni4p [125].

Localization of Chs4p homolog proteins at other locations is important for normal asexual development. S. cerevisiae Shc1p is a Chs4p homolog that is expressed specifically during sporulation. Loss of SHC1 results in decreased spore chitosan content, suggesting that Shc1p may be the Chs4p homolog that interacts with Chs3p to mediate synthesis of spore wall chitin that is subsequently deacetylated to chitosan [127]. Overexpression of CHS4 can compensate for deletion of SHC1, and overexpression of SHC1 can partially compensate for loss of CHS4, though with imperfect Chs3p localization [127]. Interestingly, Chs3p is seen in the prospore membrane in strains deficient in SHC1, thus Shc1p is important for Chs3p activity but not localization, in contrast to the importance of Chs4p for both processes at the bud neck [128]. Phosphorylation also appears to play a role in the regulation of chitin synthesis during spore formation. Loss of Sps1p, a sporulation-specific kinase, leads to incorrect localization of Chs3p and Shc1p at the prospore membrane [128]. Interestingly, Sps1p and Chs3p co-localize in cytoplasmic puncta and at the prospore membrane, though it is not known if Sps1p directly acts on Chs3p [128].

Localization of chitin synthases, both in terms of maintenance in intracellular vesicles and subsequently at specific plasma membrane locations, appears to be affected by multiple regulatory mechanisms. Transit through the secretory machinery requires the presence of multiple specialized proteins [104, 108, 111, 112, 122]. After successful navigation of the secretory pathway, establishment of activity and maintenance at the appropriate membrane location depends on the presence of a complex of various proteins, as demonstrated by the dependence of Chs3p localization at the bud neck on the presence of Bni4p and Chs4p [122, 124]. Finally, emerging research implicates phospho-signaling in the regulation of the chitin synthase regulatory proteins as well as of chitin synthases, though the precise characterization of what step in localization is altered by changes in the phosphorylation status remain to be identified [106, 107, 116, 128]. In addition, Chs3p is the only chitin synthase to have been studied in this degree of detail thus it is not clear if the regulatory mechanisms described for this particular protein are relevant to other chitin synthases.

Some version of this regulated localization exists in other pathogenic fungi, as mutations in both the C. neoformans class IV chitin synthase gene CHS3 and a chitin synthase regulator homolog CSR2 possess similar significant growth defects [49]. The level of chitosan was markedly reduced in these mutants, raising the possibility that a chitin deacetylase may directly interact with this chitin synthase [49].

It is not known if chitin synthase secretion is similarly regulated in other pathogenic yeast and nothing is known about the possibility of localization regulation by activator proteins in filamentous fungi. There are putative proteins with relatively high homology to yeast Chs5p and Chs7p in both C. neoformans and A. fumigatus (Table 2 and Table 3) [129, 130]. Interestingly, in A. fumigatus, the gene (Afu8g05620) that encodes the putative chitin synthase activator protein with the highest identity to C. albicans and S. cerevisiae Chs4p and C. neoformans Csr2 (31%, 30% and 49% respectively), is located immediately next to the gene encoding the A. fumigatus class IV chitin synthase ChsF in a head-to-head configuration, suggesting the possibility of coordinated regulation [131].

Table 2.

Percent identity ofA. fumigatus putative chitin synthase secretory and localization proteins compared to C. albicans and S. cerevisiae proteins

A. fumigatus	C. albicans		S. cerevisiae
	Protein	Identity	Protein	Identity
Afu6g02510	Chs5p	35%	Chs5p	39%
Afu3g12800	Chs6p	19%	Chs6p	20%
Afu1g12040	Chs7p	49%	Chs7p	45%
Afu3g05580	Chs4p	12%	Chs4p	12%
Afu6g02940	Chs4p	27%	Chs4p	28%
Afu8g05620	Chs4p	31%	Chs4p	30%
Afu1g16680	Bni4p	18%	Bni4p	16%

Table 3.

Percent identity of C. neoformans putative chitin synthase secretory and localization proteins compared to C. albicans and S. cerevisiae proteins

C. neoformans	C. albicans		S. cerevisiae
	Protein	Identity	Protein	Protein
CNAG_04321	Chs5p	23%	Chs5p	21%
CNAG_03325	Chs6p	18%	Chs6p	19%
CNAG_06898	Chs7p	39%	Chs7p	52%
Csr1	Chs4p	27%	Chs4p	49%
Csr2	Chs4p	38%	Chs4p	41%
Csr3	Chs4p	20%	Chs4p	26%
CNAG_02071	Bni4p	13%	Bni4p	13%

Primary septal localization of yeast Chs2p is tied to the cell cycle

Though they contribute only a minor portion of the overall cellular chitin content, the class II yeast chitin synthases are functionally important with a major role in primary septum formation. These proteins localize in positions and at times that are consistent with a role in septation, both in yeast and filamentous fungi [12, 101, 132, 133]. Studies in S. cerevisiae suggest class II synthase proteins are rapidly degraded and are closely tied to the cell cycle, presumably to prevent ectopic synthesis of primary septa [101, 102]. Chs2p remains in the ER until mitotic exit occurs, and is then transported to the membrane via Sec12p-dependent mechanisms [134]. The protein is found at the bud neck at the end of mitosis and provides structural support, possibly via chitin synthesis, that helps to maintain the stability of actin-myosin ring during constriction [101, 102]. As with Chs3p, phosphorylation is important for localization, though in the case of Chs2p phosphorylation appears to prevent rather than promote plasma membrane localization. Mutation of four putative cyclin dependent kinase consensus sequence sites in Chs2p to mimic constitutive phosphorylation or dephosphorylation either leads to retention of Chs2p in the ER even after exit from mitosis or an uncontrolled exit from the ER, respectively [93, 135].

Regulation of localization of filamentous fungal chitin synthases depends on the cytoskeleton

Studies of the regulation of other classes of chitin synthases, particularly in filamentous fungi, have been more descriptive. Consequently, little is defined mechanistically. Many of the chitin synthases, irrespective of class, are variably seen at septa, tips, and reproductive structures confirming the importance of chitin synthesis in the structural integrity of the cell wall at a variety of locations and a variety of developmental stages [12, 58, 132, 133]. Proteins have also been seen in cytoplasmic patches suggestive of chitosomes and vacuoles, suggesting that controlled secretion along with possible recycling or degradation may be common regulatory motifs, though the regulatory proteins may not generally be shared.

Discovery of the class V and VII chitin synthases, specifically found in filamentous fungi, raised interesting questions regarding the role of specific chitin synthases in the promotion of hyphal growth, particularly given the generally more dramatic growth defects seen in these gene class mutants. In addition, the combination of an amino terminal myosin motor domain and a carboxy terminal chitin synthase domain suggested the possibility of physical interaction with the cytoskeleton as a mechanism for retention of these enzymes in specific membrane locations. Studies in A. nidulans and W. dermatitidis have demonstrated actin binding by the myosin motor domain, and dependence on an intact actin network and actin binding for localization as well as functionality [136–138]. In the plant fungal pathogen Ustilago maydis, studies have differentiated between the interactions important for delivery of chitin synthases to the hyphal tip and those needed for localization to the apical plasma membrane [139]. U. maydis class V chitin synthase gene mcs1 depends on microtubules but not actin or an intact myosin motor domain for long-range vesicular movement, but intact actin and a functional myosin motor domain is needed for plasma membrane localization [139].

Conclusions and future directions

Fungal chitin synthesis is a complex and highly regulated process. Whereas transcription appears to be affected by a variety of external and internal factors, these changes in mRNA levels do not necessarily correlate with changes in chitin synthase activity or cell wall chitin content. Multiple signaling pathways have been implicated in chitin synthase regulation, at both transcriptional and post-transcriptional levels. Recent research has suggested that localization of chitin synthase localization likely represents a major regulatory mechanism. Studies with the class IV chitin synthase, S. cerevisiae Chs3p, suggest that both transport through a specialized secretory apparatus and protein localization to submembrane domains may be regulated. To date, regulation of localization of other chitin synthases has not been studied in equal detail. At this point, much of the regulatory machinery is not necessarily shared among different chitin synthases, thus it will be difficult to simultaneously target multiple proteins with a single approach. An in depth understanding of the precise roles of each protein in cell wall maintenance and repair will be essential to identifying the most likely therapeutic targets.

Simultaneous targeting of multiple cellular pathways within the same pathogen is an accepted method to maximize therapeutic efficacy. Inhibition of a single fungal cell wall component has been shown to be effective for certain fungal infections, but cell wall compensatory responses may limit efficacy. To date, chitin synthase inhibitors have not been successfully employed clinically [16, 140]. Regulation of chitin synthesis occurs on multiple levels, thus targeting of these pathways may represent an alternative approach to achieving chitin synthesis inhibition.

Figure 4.