Local Fungal Endophytes as Rich Sources of Chitinase...

Keywords: Aspergillus tubingensis, chitinase, Daldinia eschscholzii, fungal endophyte, GH18

*Corresponding Author: [email protected]

Local Fungal Endophytes as Rich Sources of Chitinase Genes

Philippine Journal of Science148 (3): 575-582, September 2019ISSN 0031 - 7683Date Received: 01 Feb 2019

Zabrina Bernice L. Malto1, Christine Jurene O. Bacal1,2,Mark Jeffrey S. Diaz1, and Eizadora T. Yu1*

1Institute of Chemistry, College of Science, University of the PhilippinesDiliman, Quezon City 1101 Philippines

2Natural Sciences Research Institute, University of the PhilippinesDiliman, Quezon City 1101 Philippines

The ability of three fungal endophytes (JB10, JB11, and D12 isolates) to degrade chitin, and their potential as microbial sources of chitinases was investigated. Amplification and sequencing of the ITS regions revealed the identity of the fungal isolates: JB10 (Fomitopsis sp.), JB11 (Aspergillus tubingensis), and D12 (Daldinia eschscholzii). All three fungi were able to grow on minimal media with colloidal chitin as sole carbon source, albeit at different rates. Isolates JB11 and D12 are observed to have comparable or faster growth rates in chitin as compared to the simpler potato dextrose carbon source. Turbidimetric measurements show that the fungal cultures are able to degrade chitin with 3–5 d of incubation. While the crude, secreted proteins from these three fungi show comparable total chitinolytic activities (~0.35 U/mL), JB11 was found to have the highest exochitinase activity (~0.25 U/mL). Bioinformatic analysis of the chitinase (GH18) genes for A. tubingensis (JB11) and D. eschscholzii (D12) revealed variability in the GH18 chitinase sequences in terms of the amino acid sequences of the canonical DXXDXDXE catalytic motif as well as the presence of additional domain architectures, which make these fungi ideal sources for chitinases for both biotechnology applications and chitinase enzyme mechanistic studies.

INTRODUCTIONThe processing of crustacean products generates a lot of shell waste as crustacean meat accounts for less than 50% of the animal’s body mass. Efforts to valorize and refine these crustacean shells for chemicals (e.g., amino acids, calcium carbonate, and N-acetyl glucosamine or NAG) are being explored to create a high-value supply chain for what is normally just discarded as waste (Yan and Chen 2015). While the complete breakdown of chitin to NAG is desirable, depolymerization to lower molecular weight chitin oligosaccharides (CTOS) can also yield high-value products with biomedical applications. Enzymes such as chitinases can degrade chitin into CTOS or NAG monomers by hydrolyzing

the glycosidic bonds under physiological conditions. These chitin-degrading enzymes can be endochitinases, which cleave the polymer internally producing CTOS or exochitinases, which cleave the reducing end of the polymer producing NAG [for N-acetyl glucosaminidases (NAGase)] or (NAG)2 (for chitobiosidases) (Hamid 2013). These enzymes can, therefore, be used in the processing of crustacean waste to produce CTOS or NAG precursors.

Microorganisms such as fungi have been used as sources and producers of economically important enzymes (Lange et al. 2012). Fungi express glycoside hydrolase family 18 (GH18) chitinases, which are primarily needed for their growth and morphogenesis (Seidl 2008, Huang et al. 2011). Recent explorations in sequencing full fungal genomes reveal the presence of multiple GH18 genes and

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the variability in protein domain structures, showing the great potential of mining chitinases from fungi (Karlsson and Stenlid 2008, Hartl et al. 2012). Several fungal chitinases have been isolated, purified or recombinantly expressed, and enzymatically characterized (Ike et al. 2006, Brzezinska and Jankiewicz 2012, van Munster et al. 2012). Discovery of novel and more effective chitinases can be increased further by studying ectoenzymes of endophytic fungi thriving in unique environments. Because of the unique habitats and the diversity of plants that have yet to be studied in the Philippines, fungal endophytes isolated from local sources could be an interesting, untapped natural source of chitinases.

One route to identifying active fungal chitinases is to fractionate the secretome proteins, screen for enzymatic activity, and identify proteins by mass spectrometry. However, this route may be laborious at times, requiring large concentrations of secretome proteins for downstream enzyme activity and MS experiments. Another way to identify and prioritize potential enzymes is to mine the genome for chitinase genes and analyze the protein sequences, thereby prioritizing genes for future recombinant protein expression and activity screening. The latter route is especially advantageous if the genomes of the target organisms of interests have been sequenced and annotated.

In this study, we screened several fungal isolates for their ability to degrade chitin and selected a few candidates as potential sources of chitinases. Standard spectroscopy-based assays were used to characterize the chitinolytic activity of the secreted proteins from three fungal candidates. Sequence analysis of chitinase genes derived from the fungal genomes was performed and revealed the presence of several chitinases with varying protein sequences and domain architectures.

MATERIALS AND METHODS

Preparation of Colloidal ChitinColloidal chitin was prepared from shrimp shells (Murthy and Bleakley 2012). Shrimp shells with heads removed were washed thoroughly, air-dried, and pulverized using a grinder. Twenty grams (20 g) of powder was demineralized by mixing 100 mL of 1 M hydrochloric acid solution for 24 h at room temperature (25 ± 3 °C). The demineralized powder was rinsed three times with deionized water and was deproteinized by incubating in 20 mL of 0.5% (w/v) of sodium hydroxide for 30 min with constant stirring at 95 °C. The resulting solid was rinsed several times with deionized water. Concentrated hydrochloric acid (150 mL) was added to the chitin and

was subjected to constant stirring for 1 h. The resulting mixture was filtered through eight layers of cheesecloth; 100 mL of filtrate was mixed with 2 L of ice-cold distilled water and then incubated overnight at 4 °C. To collect colloidal chitin, the mixture was vacuum filtered through two layers of Whatman® filter paper 3 or, alternatively, centrifuged at 8,000 × g for 15 min. The resulting chitin cake was washed using distilled water until the filtrate or supernatant was approximately pH 7. The chitin cake was then autoclaved at 121 °C for 15 min and then stored at 4 °C until further use.

Fungal Culture ConditionsFungal cultures were grown on PDA (39 g/L) at room temperature (25 ± 3 °C) and passaged every 7 d. For investigating fungal growth in chitin, fungal isolates were grown on agar plates containing M9 media supplemented with 2 mM MgSO4, 0.1 mM CaCl2, trace elements, and 2% w/v colloidal chitin). PDA, chitin, and control agar plates were inoculated with ~ 5 mm fungal mycelium plugs (placed in the center of the plate) and allowed to grow at room temperature. Radial growth measurements were taken daily for 10 d, marking points along the circumference of fungal mycelia, until growth reached the edge of the plate. Growth measurements were taken from at least three replicate plates. Radial growth rate (Kr) was estimated using colony diameter versus time during constant growth (Reeslev and Kjøller 1995) (Table 1). The fungal strains used in this study are Daldinia sp. D12 isolated from bamboo (Bambuseae sp.), Fomitopsis sp. JB10, and Aspergillus tubingensis JB11 (Bacal and Yu 2017).

Table 1. Radial growth rates of fungal isolates (Kr ) grown in PDA, agar, and agar supplemented with colloidal chitin plates. The maximum diameter of fungal mycelia after 10 d growth is reported (in parenthesis). The plates used were 80 mm diameter plates.

Fungal IsolatesKr

a (mm/d)

PDA Agar Chitin

JB10 11.3(80 mm)

2(17 mm)

2.7(23 mm)

JB11 3.95(39 mm)

6.4(40 mm)

8.7(72 mm)

D12 33.5(80 mm)

0.85(15.5 mm)

17.7( 80 mm)

aKr was estimated using colony diameter versus time (~ Day 2–5).

Liquid Culture Conditions and Semi-quantitative Chitinase Activity MonitoringIsolates were initially grown in 50 mL of potato dextrose broth (PDB) for 5 d. Since the standardization of mycelium-based inoculum is difficult (Fazenda et al.

Malto et al.: Chitinases from Fungal EndophytesPhilippine Journal of ScienceVol. 148 No. 3, September 2019

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2011), mycelia on top of ~ 1 mm x 1 mm plugs were allowed to grow directly in PDB. Standardization was based on the method of El-Hadi et al. (2014) with slight modifications, wherein 2 mL of the inocula was pipetted out from the PDB culture and transferred to the basal media for chitinase production containing 10 g/L colloidal chitin, 0.5 g/L ammonium sulfate, 10.0 g/L potassium dihydrogen phosphate, 0.1 g/L magnesium sulfate heptahydrate, and 0.2 g/L sodium chloride. The cultures were placed in a rotary shaker for discontinuous shaking (100 rpm) at ambient temperature (25 ± 3 °C) for 7 d. Semi-quantitative monitoring of chitinase activity was done by comparing the absorbance at 510 nm of the medium containing the isolate and the control containing the medium only (Harman et al. 1993). A reduction of turbidity of the liquid culture medium can be attributed to the formation of more soluble chitin oligomers during chitin breakdown (Harman et al. 1993).

Quantitative Enzyme Activity AssaysCrude secreted enzyme extracts were acquired after 7 d by filtering the culture media using two layers of Whatman® 3 filter paper and then centrifuged at 8,000 × g for 30 min. The supernatant (50 mL) was collected and the proteins were concentrated 50 times via ultrafiltration using Amicon® stirred cell (Millipore, MWCO 10 kDa). The concentrated proteins were stored at 4 °C until use for chitinase activity determination.

Total Chitinase ActivityTotal chitinase activity of the crude extracts was determined through a microplate-based dinitrosalicylic acid (DNS) assay (King et al. 2009). Substrate solution was prepared by partially dissolving colloidal chitin (1.25% w/v) in 200 mM potassium phosphate buffer pH 6.0 in deionized water. The solution (100 μL) was mixed with the same volume of the concentrated extract and then incubated for 2 h at 50 °C in shaking water bath. The reaction was stopped by putting the solutions in the freezer (–20 °C) for 5 min. Unreacted colloidal chitin was removed through centrifugation (10,000 × g for 10 min). DNS reagent (80 μL) was added to 40 μL of the solution and heated at 95 °C for 5 min. The absorbance of the resulting solution was measured at 540 nm. Enzyme controls, substrate control, and reagent blank were also prepared. Standard solutions for the calibration curve were prepared using NAG with concentration ranging 50–800 μg/mL by dissolving NAG to 200 mM potassium phosphate buffer pH 6.0. Standard solution (40 μL) was mixed with 80 μL of DNS reagent, heated at 95 °C for 5 min, and read at 540 nm. One unit of enzyme activity (U) using colloidal chitin as substrate was defined as the liberation of 1 μmol of NAG.

NAGase Activity AssayNAGase activity was determined using a standard assay adapted from Liu et al. (2013). The assay was performed in a 96-well plate using 4-nitrophenyl N-acetyl-β-D-glucosaminide (pNP-NAG) (Sigma-Aldrich, N9376) as substrate. The crude enzyme extract (10 μL, ~0.096 μg protein) was mixed with 90 μL of substrate solution (1 mg/mL in 50 mM acetate buffer, pH 5) and then incubated for 10 min at 50 °C. Control assays without crude enzymes (90 μL of 1 mg/mL solution of pNP-NAG with 10 μL of reaction buffer) and without the substrate (10 μL of the extract with 90 μL of 100 mM acetate buffer, pH 5) were also prepared. Blank solutions contained 100 μL of buffer only. Two hundred microliters (200 μL) of stop solution containing 1M sodium carbonate was immediately added to each reaction well after 10 min of incubation. Standard solutions for the calibration curve were prepared using p-nitrophenol (2–100 μg/mL). The absorbance measurements of the resulting solutions were read at 405 nm. One unit of enzyme activity (U) was defined as the release of 1 μmol of p-nitrophenol.

Sequence Analysis of Chitinase GenesAnnotated chitinase genes (GH18) of Daldinia eschscholzii and Aspergillus tubingensis were obtained from JGI Mycocosm website (Nordberg et al. 2014, Wu et al. 2017, de Vries et al. 2017). Multiple sequence alignment of the chitinase genes within and between genomes was performed using MEGA7 by using the maximum likelihood model with 500 bootstrap iterations (Kumar et al. 2016). The protein domains were determined using InterPro and Pfam (Finn et al. 2016, 2017).

RESULTS Previous work in our lab showed that several fungal isolates secreted glycolytic enzymes and yielded high enzymatic indices when screened for cellulose degradation (Bacal and Yu 2017). In nature, these endophytes obtain nutrition from the host plant they colonize, thus should have the necessary biochemical machinery to obtain sugars from cellulosic biomass. Due to the similarity of the glycosidic linkages in cellulose and chitin, isolates that yielded high enzymatic indices (⪴ 1) in the previous study were chosen for chitinase activity screening. We wanted to determine whether these fungal isolates can use chitin and followed the growth of these fungal isolates on potato dextrose and M9 salts supplemented with colloidal chitin (Table 1).

All three isolates were able to grow on chitin, albeit at different rates. Consistent with previous growth monitoring studies, JB10 preferred the potato dextrose


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media over the more complex sugar sources chitin and carboxymethylcellulose (CMC) (Bacal and Yu 2017). Interestingly, JB11 grew much faster in colloidal chitin as compared to potato dextrose and agar control. Both potato dextrose and colloidal chitin supported the growth of D12, with the D12 mycelia able to grow and fill the whole diameter of the plates within four to six days. This observation suggests the non-selective nature of D12 for carbon sources as it is able to utilize both simple and complex carbohydrates to support its growth.

Next, the ability of these fungal endophytes to depolymerize chitin was measured using the modified assay by Harman et al. (1993). Figure 1 shows the results of the monitoring of chitin breakdown. It is noticeable that isolates JB10 and JB11 were able to efficiently break down the colloidal chitin within a few days after inoculation. Meanwhile, D12 was able to effectively depolymerize the colloidal chitin only after five days post-inoculation. The negative control (no fungi added) remained turbid throughout the duration of the experiments. The genomes of two of the fungal endophytes (JB11 and

D12) have been sequenced and are publicly available (Nordberg et al. 2014, de Vries et al. 2017, Wu et al. 2017). We then sought to mine the Aspergillus tubingensis (JB11) and Daldinia eschscholzii (D12) genomes for annotated GH18 genes. Multiple sequence alignment of the amino acid sequences of the GH18 genes was performed to determine the sequence similarity of the GH18 genes within the fungal isolate and with GH18 from other related fungi.

There are 16 GH18 genes in the A. tubingensis genome and these genes have 72–100% sequence identity with GH18 gene sequences from other fungi belonging to the order Eurotiales. The A. tubingensis GH18 genes clustered into three main groups showing differences in protein domain (Figure 3). There is a high degree of conservation of the canonical catalytic DXXDXDXE motif domain for chitinases (Figure 3C), with only two predicted GH18 genes without this catalytic domain (Asptu1 130331 and Asptu1 61666) (Lu et al. 2002, Huang et al. 2011). Despite the lack of the chitinase DXXDXDXE motif, these genes were annotated as GH18 probably due to some sequence similarity to the chitinase motif as well as having a chitinase insertion domain in the sequences. Interestingly, the majority of the GH18 sequences contain signal peptides, indicating that the protein products are targeted for secretion (Huang et al. 2011). Based on the GO gene annotation, only two GH18 gene products (Asptu1 18926 and Asptu1 191924) appear to be membrane-bound. In addition to the chitinase catalytic domain, other GH18 genes appear to have additional domains that may contribute to its efficiency as an enzyme. Seven of the GH18 genes have additional known substrate-binding domains such as the chitin-binding type I domain and

Figure 1. Chitin depolymerization by fungi as monitored by turbidimetry measurements. Fungal endophytes were grown in liquid culture containing colloidal chitin as the sole carbon source. Turbidity of the medium was measured by reading the absorbance at 510 nm. Data are presented as % reduction in turbidity vs. the control containing the medium only.

Figure 2. Chitinase activity of crude fungal protein secretomes. Total chitinase activity was measured using the DNS assay using colloidal chitin as substrate. The NAGase activity was measured using the pNP-NAG assay. One unit of activity is equal to 1 µmol reducing sugar (NAG), and 1 μmol of p-nitrophenol measured per min per mL of protein extracts, respectively.

The chitinolytic activity of the secreted extracellular fungal enzymes was determined using standard spectroscopic assays. Figure 2 shows a summary of the chitinolytic activity of the secreted fungal proteins. The total chitinolytic activity was determined using the DNS assay and colloidal chitin as the substrate. The crude protein mixtures from all isolates show comparable total chitinase activities. In order to further define the contribution of the exochitinases present in the protein mixture, we used the same protein mixtures and subjected them to a more specific pNP-NAG assay. Figure 2 shows that the isolate JB11 exhibited the highest exochitinase activity among the three isolates.


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the LysM domains. These domains aid in the hydrolysis of insoluble chitin by providing a binding site for the extracellular polysaccharides (Buist et al. 2008, Huang et al. 2011). Asptu1 124210 contains a domain that codes for an Ecp2 effector protein that is implicated as a necrosis-inducing factor in plants and contributes to fungal pathogenicity (Stergiopoulos et al. 2010).

On the other hand, we found 18 annotated GH18 genes in the D. eschscholzii (D12 isolate) genome. These GH18 genes appear to be unique as they only have around 18–77% sequence with annotated GH18 gene sequences from the class Sordariomycetes. Figure 4 shows the sequence alignment, homology and domain architecture for the D. eschscholzii GH18 genes. Although all of the Daldinia GH18 genes contain the catalytic domain, close inspection of the sequences shows more amino acid differences, even for some of the highly conserved aspartic residues that make up the DXXDXDXE motif (Figure 4C). Twelve out of the 18 chitinase genes contain signal peptides, signifying that these are probably secreted proteins. However, unlike in A. tubingensis, only a few GH18 genes contain additional substrate-binding domains.

DISCUSSIONThe availability of local sources of microbial enzymes will benefit industries that use and import enzymes for biotechnological applications. Discovery of local microbial sources of enzymes is necessary to support Philippine-based enzyme biotech endeavors. In this study, we focused on the discovery and identification of promising sources of fungal chitinases as we performed

bioinformatic analysis and enzyme characterization of the secreted chitinolytic enzymes. As a first approximation, the ability of fungi to grow with chitin as the sole carbon source depends on the isolate’s ability to express efficient chitinases, which degrade chitin into lower molecular weight CTOS that they can then acquire as direct nutrient sources. Three fungal endophytes – JB10, JB11, and D12 – isolated from different plant sources have the ability to grow using colloidal chitin as sole carbon source, albeit at different rates.

It is interesting to note from the fungal studies, that these fungi appear to have different preferences in terms of the complexity of carbon source. Typical fungi are easier to grow in media using simpler carbohydrates (e.g., potato dextrose); however, JB11 appears to favor the more complex carbohydrates (Liu et al. 2013). The relatively fast growth rates of D12 in solid agar – either potato dextrose, chitin, or CMC (data not shown) – reveal that D12 is not as selective as typical fungi. The growth rate measurements are supported by turbidimetric analysis of the chitin substrates as well as enzymatic assays, showing that the secreted fungal protein extracts have chitinolytic activity.

The potent glycoside hydrolase activities of enzymes from Aspergillus fungi are well documented. In fact, quite a few of recombinant enzymes originate from this family of fungi (Brzezinska and Jankiewicz 2012, van Munster et al. 2012). Due to the high exochitinase activity exhibited by JB11 secreted proteins, isolation and further characterization of the JB11 exochitinase should and will be pursued. Alternatively, expression of the A. tubingensis chitinase genes can be performed and it will be interesting to note if the presence of the additional chitin-binding domains will increase the efficiency of the chitinases.

Figure 3. Protein sequence alignment and domain architecture of A. tubingensis GH18 genes. (A) Multiple sequence alignment of the amino acid sequences of A. tubingensis (JB11) chitinases using the maximum likelihood method in MEGA 7 with 500 bootstrap iterations and a WAG+G substitution mode. (B) Modular domain architecture of the chitinase genes showing the presence and relative locations of the canonical catalytic motif (red), chitin-binding type 1 (green), LysM domain (blue), and an Ecp2 effector protein domain (yellow). (C) Sequence conservation of the catalytic chitinase domain using BlockLogo (Olsen et al. 2013).


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In contrast, there is not much known about chitinases from Daldinia species. While bioinformatic analysis has been done on chit genes of other Daldinia isolates (Chan et al. 2015), our biochemical study provides a first approximation of the chitinolytic activity of secreted proteins from a Daldinia fungus. Bioinformatic analysis of the chit genes from D. eschscholzii also reveals more variation in the chitinase catalytic domain sequence, with sequence variants for the highly conserved DXXDXDXE consensus sequence. Expression of these chitinases would allow mechanistic studies on the effect of the sequence variants on the catalytic efficiency of the Daldinia chitinases.

CONCLUSIONWe report the isolation of fungal endophytes and the determination of the chitinolytic activity of their secreted proteins. Bioinformatic analysis of the GH18 genes from A. tubingensis and D. eschscholzii reveals that majority contain signal peptides and contain additional substrate binding motifs that may aid in their efficiency to break down chitin. Future biochemical characterization on recombinantly expressed chitinases is necessary to link the chitinolytic activity of the secreted fungal proteins with the annotated GH18 genes from these fungi. Lastly, the variability of GH18 chitinase catalytic domain sequences from JB11 and D12 makes them ideal sources of chitinases for both biotechnology applications and chitinase enzyme mechanistic studies.

ACKNOWLEDGMENTSThis project was funded by the UP Natural Sciences Research Institute (NSRI CHE-16-1-05) and in part by a UP OVPAA Balik PhD grant to E. Yu. Plant samples were identified by UP Diliman Institute of Biology Herbarium.

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