Genome-wide identification and characterization of Chitinase gene family in Brassica juncea and Camelina sativa in response to Alternaria brassicae

Genomics xxx (xxxx) xxx–xxx

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Original Article

Genome-wide identification and characterization of Chitinase gene family in Brassica juncea and Camelina sativa in response to Alternaria brassicae Zahoor Ahmad Mira,e,1, Sajad Alia,b,1, S.M. Shivarajc, Javaid Akhter Bhatd, Apekshita Singhe, ⁎ Prashant Yadava, Sandhya Rawata, Pradeep K. Paplaof, Anita Grovera, a

National Research Centre on Plant Biotechnology, NRCPB, New Delhi, India Centre of Research for Development, University of Kashmir, Srinagar, India c Laval University, Quebec, Canada d State Key Laboratory for Crop Genetics and Germplasm Enhancement, Nanjing Agricultural University, Nanjing 210095, China e Amity Institute of Biotechnology, Amity University Noida, India f Division of Nematology, IARI, New Delhi, India b

A R T I C LE I N FO

A B S T R A C T

Keywords: Brassica juncea Camelina sativa Alternaria brassicae Pathogenesis related genes Brassicaceae Chitinases

Chitinases belong to the group of Pathogenesis-related (PR) proteins that provides protection against fungal pathogens. This study presents the, genome-wide identification and characterization of chitinase gene family in two important oilseed crops B. juncea and C. sativa belonging to family Brassicaceae. We have identified 47 and 79 chitinase genes in the genomes of B. juncea and C. sativa, respectively. Phylogenetic analysis of chitinases in both the species revealed four distinct sub-groups, representing different classes of chitinases (I-V). Microscopic and biochemical study reveals the role of reactive oxygen species (ROS) scavenging enzymes in disease resistance of B. juncea and C. sativa. Furthermore, qRT-PCR analysis showed that expression of chitinases in both B. juncea and C. sativa was significantly induced after Alternaria brassicae infection. However, the fold change in chitinase gene expression was considerably higher in C. sativa compared to B. juncea, which further proves their role in C. sativa disease resistance to A. brassicae. This study provides comprehensive analysis on chitinase gene family in B. juncea and C. sativa and in future may serve as a potential candidate for improving disease resistance in B. juncea through transgenic approach.

1. Introduction Plants being sessile are under robust selection to respond adaptively to environmental stress, among which the biotic stresses are the major one causing significant losses in crop yield [1]. In this regard, plants have evolved an array of mechanical and chemical defense mechanism to overcome abnormalities caused by these stresses. Plants combat biotic stresses by synthesizing several compounds such as pathogenesisrelated (PR) proteins that constitute a small group of heterogeneous proteins [2]. The PR proteins are specific proteins that play a key role in plant disease resistance and are usually induced by pathological conditions [3]. In addition to biotic stresses, these proteins are also induced in response to abiotic stresses; hence are generally considered to be part of multiple defense system in plants [4]. PR proteins are classified into 17 families with diverse functions; some of these are PR1 (unknown), PR2 (β-1, 3-glucanase), PR3 (chitinases), PR5 (thaumatin like), PR9

(peroxidases), PR12 (plant defensins) and PR13 (thionins) [5]. PR3/ Chitinases, are a group of PR proteins that catalyses the hydrolysis of the chitin present in fungal cell wall, thereby inhibiting the fungal pathogens [6]. These proteins are reported to show basal level of expression under normal conditions, but following infection their expression increases dramatically at the site of infection and also results in systemic acquired resistance (SAR) [7]. Chitinases have been found in a diverse range of organisms belonging to different kingdoms of life [8,9]. According to the amino acid sequence similarity of the catalytic domains, chitinases are grouped into families 18 and 19 of glycosyl hydrolases (GH18 & GH19) [10]. Based on phylogeny, mechanisms of catalytic reactions, three-dimensional (3D) structures, sensitivity to inhibitors and other characteristics, these families are further categorized into five distinct classes (class I-V) [11]. Chitinases from GH18 family fall under classes III and V, which are found widely distributed across different organisms such as

⁎

Corresponding author. E-mail address: [email protected] (A. Grover). 1 These authors have contributed equally. https://doi.org/10.1016/j.ygeno.2019.05.011 Received 12 February 2019; Received in revised form 30 April 2019; Accepted 10 May 2019 0888-7543/ © 2019 Elsevier Inc. All rights reserved.

Please cite this article as: Zahoor Ahmad Mir, et al., Genomics, https://doi.org/10.1016/j.ygeno.2019.05.011

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retrieved from the Brassica database (BRAD) (http://brassicadb.org/ brad/). Protein sequences local database of B. juncea and C. sativa genes were created in Bioedit ver. 7.2.5 [27]. The 24 chitinase genes of A. thaliana downloaded from The Arabidopsis Information Resource (TAIR) database (https://www.arabidopsis.org/) were used as query to identify putative orthologs of chitinase genes in B. juncea and C. sativa local database using BLASTp. To identify high scoring pairs (HSPs), we kept initial cut-off for an e-value of 10−5. The HSPs showing e-value cut-off 10−5 were selected based on tabulated blast output. Finally, unique sequences were selected by removing redundant hits with highest similarity for the further analysis. The entire set of unique sequences were submitted to Pfam and SMART (http://smart.emblheidelberg.de/) to confirm the presence of the chitin binding domain (CBD). To remove sequences that lack CBD we performed multiple sequence alignment (MSA).

bacteria, fungi, animals, viruses, and higher plants. However, members of GH19 family are mainly found in higher plants and some bacteria, and belongs to class I, II, and IV chitinases. As plant do not possess chitin in their cell wall and other organelles which makes them an unusual source of chitinases. The plant chitinases catalyze hydrolysis of chitin and releases chito-oligosaccharides, which may serve as elicitors to activate plant chitinase genes in a positive feedback cycle [12]. This is a successful immune strategy adopted by plants against microbes and herbivores [11,13,15]. Therefore, chitinases are classified as PR proteins and are considered important targets for crop improvement [2]. Brassica juncea (Indian mustard) and C. sativa are the two important oilseed crops that belong to family Brassicaceae and are an essential source of edible oil worldwide. However, there are various biotic and abiotic stresses that decreases the crop productivity of above crops, but the Alternaria blight is reported to be the major constraint caused by A. brassicae [19]. Alternaria blight appears every year (endemic) and causes up to 36.88% loss of yield in mustard [20]. However, yield losses in C. sativa due to Alternaria blight is considerably lower compared to B. juncea, because former have been reported to show increased resistance to this disease [21,70]. It has been widely confirmed that C. sativa is resistant to A. brassicae and Alternaria brassicicola (related pathogens) [22], hence can be potentially exploited in the development of resistant Brassica varieties. To develop pathogen-resistant varieties it is important to mine key genes imparting resistance. In almost all plant species the chitinase family members have been identified, and in vitro studies has revealed that many of these genes inhibit fungal growth [23], for example overexpression of chitinase genes from plant and fungal origin have enhanced the resistance to fungal pathogens in a different transgenic plant [24]. However, till date, chitinase genes have not been systematically analyzed in B. juncea and C. sativa. Although, the B. juncea lines have been transformed with some chitinases from other organisms, but the resistance of these transgenic lines to pathogens is not clear [25]. The oxidative burst caused due to the biotic stress which leads to the production of ROS is a complex process in plants and it activates the mitogen activated protein kinases and provides resistance against invading pathogen [70,71]. Necrotroph's are assumed to cause cell death in plants and get benefited from the ROS produced by the plants [72] and the signal perceived is carried out to cells leads to the production of hydrogen peroxide (H2O2) which further activates the ROS scavenging enzymes to neutralize the fungal toxins. Defense enzymes such as SOD (superoxide dismutase), CAT (catalase), POX (peroxidase) and glutathione reductase, as well as some non-enzymatic antioxidants such as glutathione (GSH), carotenoids, tocopherols, and other phenolic compounds etc. [22], as it has been demonstrated that during environmental stresses activity of defense-related enzymes increase several folds [73]. With availability of sequence data of plant genomes and transcriptomes, the genome-wide identification of gene families becomes easily accessible [26]. In this regard, the availability of whole-genome data of B. juncea and C. sativa offers an opportunity to identify chitinase genes on a genome-wide scale. Consequently, in the present study genome-wide identification of chitinase gene family was performed in B. juncea and C. sativa. Furthermore, the expression profiling of chitinase genes of B. juncea and C. sativa under biotic stress (A. brassicae infection) were analyzed using qRT-PCR. Hence, the results of the present study provide first comprehensive classification of chitinases in these Brassicaceae species and highlighted the importance as well as possible roles of chitinases in biotic stress alleviation.

2.2. Phylogenetic analysis and multiple sequence alignment The CLUSTALW alignment function in MEGA 7.0 [28], were used to align chitinase sequences. Maximum likelihood method was used to construct the phylogenetic tree, and 1000 bootstraps were used to measure the stability of the branch node. The chitinases were grouped in accordance with different chitinase classes (I-V) of A. thaliana. The phylogenetic tree was constructed using B. juncea and C. sativa chitinase sequences and two separate trees were constructed using B. juncea and C. sativa chitinase sequences, respectively with the common 24 chitinase sequences of Arabidopsis. 2.3. Gene structure and chromosomal localization The gene and CDS sequence of chitinases identified for B. juncea and C. sativa genome were retrieved from BRAD database (http:// brassicadb.org/brad/). The exon-intron structures of chitinase encoding genes of both crop species were determined based on alignment of CDS sequences with corresponding genomic sequences, and graphical display was created using the online Gene Structure Display Server (2.071) (http://gsds.cbi.pku.edu.cn/). The positions of the putative chitinase genes were visualized by mapping to the 18 and 20 chromosomes of B. juncea and C. sativa, respectively using base pair start positions in Mapchart2.2 [29]. 2.4. Distribution of conserved motifs and 3D structure of Chis genes The upstream region (1.5 kb) of randomly selected five chitinase genes each from B. juncea and C. sativa were analyzed for conserved cisregulatory elements involved in multiple stresses using Plant Care Database (http://bioinformatics.psb.ugent.be/webtools/plantcare/ html/). For final quality check three dimensional (3D) models of the same chitinase proteins for both crop species were constructed using Phyre2 server [30]. For structural models the quality control and thresholds are as follows: alignment coverage > 65% and confidence = 100%. MEMSAT-SVM prediction method available in Phyre2 server was used to predict transmembrane helix and topology of the chitinases. 2.5. Genome Synteny and gene duplication The syntenic information of A. thaliana, B. rapa, B. oleracea and C. sativa was downloaded from the BRAD database (http://brassicadb. org/brad/), whereas the same information is not available for B. juncea and therefore it is not included in synteny analysis. Chitinase genes were mapped to the syntenic blocks for inter-genomic comparison. The syntenic diagram was drawn using Circos software version 0.63. Tandem duplications of chitinase genes in the C. sativa genome were identified based on their physical locations on individual chromosomes. Genes having an adjacent homologous ID gene on the same C. sativa

2. Methods 2.1. Genome-wide identification and distribution of chitinase genes in B. juncea and C. sativa The amino acid sequences of B. juncea and C. sativa genome were 2

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enzymatic reactions. Reaction was setup and 50 mM Tris-HCl buffer (pH 7.6), 1 ml NADPH (0.15 mM), 100 μl oxidized glutathione (1 mM GSSG), 3 mM MgCl2, and 0.3 ml enzyme extract was added in a 2 ml centrifuge tube. Activity of GR was determined by measuring absorbance decline of NADPH at 340 nm and enzyme activity was expressed as NADPH oxidized μmol min−1 mg−1 protein.

chromosome with no more than one intervening gene was considered to be tandemly duplicated. 2.6. Culture and inoculation with A. brassicae To examine the induction of B. juncea and C. sativa chitinase genes, 40 days old plants of both crop species were infected with A. brassicae strain. The A. brassicae culture was collected from Division of Plant Pathology, IARI, New Delhi for Indian Type Culture Collection (I.D. No. 81651), and was cultured at 22 °C for 20 days on radish dextrose agar. The conidia were taken from A. brassicae and suspended in sterile distilled water, and the muslin cloth was used for filtering, and diluted to 5 × 103 conidia/ml. Spore suspension (4–6 drops) of A. brassicae (5 × 103 spores cm−3) were inoculated on four different selected spots of the leaf surface of 40 days old B. juncea and C. sativa plants, and were then incubated in a chamber at 25 °C, with relative humidity 100%. Control plants for each treatment were treated with sterile distilled water. For RNA isolation leaf samples were harvested from control and infected plants after 6, 12, 24 and 48 h post-inoculation (hpi), and after flash frozen in liquid nitrogen were stored at −80 °C. Three different plants of both B. juncea and C. sativa were infected with A. brassicae on separate occasions to provide biological replicates for qRT-PCR analysis.

2.9. RNA isolation and qRT-PCR analysis A leaf sample of 100 mg was used for the extraction of total RNA needed of both control and treated B. juncea and C. sativa seedlings using PureLink RNA Mini Kit (Ambion Life Technologies, USA). RNA concentration and quality were measured by Nanodrop spectrophotometer (NanoDrop 2000 Thermo Scientific, Wilmington, DE). Superscript III cDNA synthesis kit according to the manufacturer's protocol (Invitrogen, Canada) was used to generate first strand cDNA from 2-μg of DNaseI-treated total RNA in 20-ul reaction volume. Primers of B. juncea and C. sativa chitinase genes as well as alpha-tubulin were designed using Oligoanalyzer software (Table S1). qRT-PCR mixture was run for 5 min at 95 °C, followed by 40 cycles of 30 s at 94 °C, 30 s at 60 °C, and 30 s at 72 °C, and this mixture contained 2 μl of cDNA, 5 μl of SYBR green qRT-PCR master mix (Takara, Japan) and 0.5 μl (10 picomol) of each primer. Each reaction was carried out in triplicates, and was replicated in three biological repeats. For internal control alpha-tubulin was used as house-keeping gene in all the experiments, and delta CT method was used to check relative expression levels of each gene [31]. Significant level of expression was considered at fold changes with p values < .05.

2.7. DAB staining (3,3′-Diaminobenzidine) Leaves of 25 days old B. juncea and C. sativa were used for DAB staining described by [74]. Leaves were stained with DAB (3,3′-Diaminobenzidine) for H2O2 production and were kept in dark for 8 h after incubation leaves were destained using bleach solution (ethanol:acetic acid:glycerol = 3:1:1) at 100 °c.

2.10. Statistical analysis Three biological replicates were used for all experiments and each was repeated three times. To determine the significant difference of chitinase gene expression in control and treated samples of B. juncea and C. sativa we used student's t-test. For comparison differences between two groups of data in all the experiments were evaluated as statistically significant (⁎p < .05) or extremely significant (⁎⁎p < .01).

2.8. Enzyme Assay's Enzyme assays were carried out in B. juncea and C. sativa leaf extracts as per the [75]. The activity of SOD was determined through its ability to prevent nitro blue tetrazolium (NBT) photochemical reaction via the method of [76]. In 5 ml of 0.1 M Sodium Phosphate buffer (pH = 7) the 1 g of leaf tissue was ground and centrifuged at 12000 rpm to get supernatant that was further used for enzyme assay's to the total reaction 1.110 ml of 50 mM phosphate buffer (pH 7.4), 0.040 ml of 1% (v/v) Triton X-100, 0.075 ml of 20 mM L-methionine, 0.075 ml of 10 mM hydroxylamine hydrochloride and 0.1 ml of 50 μM EDTA reaction was started by adding 100 μl of enzyme extract and 80 μl of riboflavin (50 μM). It was kept under illuminated W-Philips fluorescent lamps (60 μmol m−2 s−1) for 10 min and the extract of peroxidase enzyme assay was prepared by homogenizing tissues in 50 mM potassium phosphate buffer (pH 7.8) the final volume of reaction contain 3 ml of 0.25% (v/v) guaiacol in potassium phosphate buffer (pH -7) containing 10 mM hydrogen peroxide reaction was started by adding 100 ul crude enzyme extract to the reaction mixture, which was measured spectrophotometrically at 470 nm (Hitachi U 2000, Japan). The last component to be added was H2O2 and the reaction was monitored by the decrease in absorbance at 290 nm (extinction coefficient of 2.8 mM−1 cm−1) up to 5 min. The infected leaves were crushed in 50 mM Tris–NaOH buffer (pH 7.0) to prepare catalase enzyme assay. In the final volume of 3 ml assay mixture contained 50 mM H2O2, 100 mM potassium phosphate buffer (pH 7.0), and 200 μl enzyme extract. The H2O2 decomposition was followed at 240 nm (extinction coefficient of 0.036 mM−1 cm−1) by a decrease in absorbance for 5 min at 25 °C. Anderson, 1996 method was followed for the assessment of GR activity, and catalase activity was expressed as μmol of H2O2 oxidized min−1 mg−1 protein. The chilled mortar and pestle were used to homogenize the leaf samples (0.1 g) in 5 ml of 50 mM Tris-HCl buffer (pH 7.6). At a frequency of 15,000 ×g homogenate was centrifuged for 30 min at 4 °C and the supernatant obtained was further used for

3. Results 3.1. Genome-wide identification and distribution of chitinase genes in B. juncea and C. sativa The availability of whole-genome sequence of B. juncea and C. sativa provides an opportunity to carry out genome-wide identification of any gene family in these crop species, provided their homologs are identified in the related plant species. Hence, in the present study, we used 24 known A. thaliana chitinase genes as queries in BLAST searche of the local database created in Bioedit programme and identified a total of 47 and 79 genes encoding chitinases in B. juncea and C. sativa, respectively (Table S1 & S2). The chitinase homologs identified in B. juncea and C. sativa genome were named as BjChis and CsChis, respectively. Furthermore, the functional annotation of these sequences using conserved domain database (CDD) revealed the presence of either Glyco_hydro 18 or Glyco_hydro 19 domain in the predicted protein sequence which is an essential component for the hydrolysis of chitin, and hence further confirmed the function of these candidate sequences as chitinases (Tables 1 & 2). The chitinases with Glyco_hydro 18 as catalytic domain belong to either Class III or V and those with Glyco_hydro 19 belongs to Class I, II or IV of GH family. In B. juncea majority of identified chitinases (41) were found to have Glyco_hydro 19 domain and only few (6) harbored Glyco_hydro 18 domain, similarly in C. sativa 51 chitinases contained Glyco_hydro 19 domain, while remaining 28 chitinases showed Glyco_hydro 18 catalytic domain (Tables 1 & 2). 3

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Table 1 Nomenclature and physico-chemical properties of chitinase genes identified in B. juncea. S.No.

Name

Class

Glyco-hydro Group

Protein length (aa)

Mol. Wt. (Da)

PI (pH)

Instability Index

GRAVY

1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47

BjuA006108 BjuA006109 BjuA012108 BjuB006956 BjuB008588 BjuB023719 BjuO006254 BjuO012596 BjuA012109 BjuO012597 BjuA031312 BjuA031440 BjuA044968 BjuB021972 BjuB043883 BjuO012598 BjuA046780 BjuB038092 BjuA010653 BjuA010654 BjuA010656 BjuA010657 BjuA010658 BjuA017884 BjuA017886 BjuA033614 BjuA035692 BjuA042004 BjuB015897 BjuB016906 BjuB016907 BjuB016908 BjuB016909 BjuB016912 BjuB020315 BjuB027046 BjuB027047 BjuB041249 BjuB046249 BjuB046489 BjuO001980 BjuO008763 BjuO008764 BjuA020034 BjuA040662 BjuB013723 BjuB020908

I I I I I I I I I I II II II II II II III III IV IV IV IV IV IV IV IV IV IV IV IV IV IV IV IV IV IV IV IV IV IV IV IV IV V V V V

Glyco_hydro_19 Glyco_hydro_19 Chitinase_glyco_hydro_19 Glyco_hydro_19 Glyco_hydro_19 Glyco_hydro_19 Glyco_hydro_19 Glyco_hydro_19 Glyco_hydro_19 Glyco_hydro_19 Chitinase_glyco_hydro_19 Chitinase_glyco_hydro_19 Chitinase_glyco_hydro_19 Chitinase_glyco_hydro_19 Chitinase_glyco_hydro_19 Chitinase_glyco_hydro_19 Glyco_hydro_18 Glyco_hydro_18 Chitinase_glyco_hydro_19 Chitinase_glyco_hydro_19 Chitinase_glyco_hydro_19 Chitinase_glyco_hydro_19 Chitinase_glyco_hydro_19 Chitinase_glyco_hydro_19 Chitinase_glyco_hydro_19 Chitinase_glyco_hydro_19 Chitinase_glyco_hydro_19 Chitinase_glyco_hydro_19 Chitinase_glyco_hydro_19 Chitinase_glyco_hydro_19 Chitinase_glyco_hydro_19 Chitinase_glyco_hydro_19 Chitinase_glyco_hydro_19 Chitinase_glyco_hydro_19 Chitinase_glyco_hydro_19 Chitinase_glyco_hydro_19 Chitinase_glyco_hydro_19 Chitinase_glyco_hydro_19 Chitinase_glyco_hydro_19 Chitinase_glyco_hydro_19 Chitinase_glyco_hydro_19 Chitinase_glyco_hydro_19 Chitinase_glyco_hydro_19 GH18_plant_Chitinase_class_V GH18_plant_Chitinase_class_V GH18_plant_Chitinase_class_V GH18_plant_Chitinase_class_V

334 297 400 282 391 322 227 203 483 199 321 279 321 321 271 199 313 301 282 229 268 250 278 282 283 254 273 262 259 277 281 280 268 286 263 259 281 262 273 382 278 296 281 380 278 380 378

37,019.96 32,022.04 42,774.09 30,317.94 42,093.98 34,684.01 24,553.86 22,625.52 53,733.77 21,732.16 35,507.33 31,424.51 35,576.54 35,521.46 30,164.82 20,403.97 34,444.05 33,032.82 30,294.88 24,846.06 28,697.21 26,852.67 30,310.12 30,317.94 30,813.31 27,515.26 29,589.1 28,505.96 27,874.38 30,079.05 31,737.55 31,279 28,483.97 31,036.88 28,876.53 28,607.26 30,248.97 28,407.85 29,480.86 41,515.37 30,143.73 32,869.98 31,719.59 41,404.23 30,143.73 41,249 42,196.96

6.3 5.04 4.84 5.33 4.48 6.84 9.24 5.68 5.97 5.26 6.23 8.98 6.53 6.52 5.97 4.35 9 8.94 5.53 8.95 8.09 9.69 9.03 5.33 9.44 9.1 4.85 4.93 7.82 9.03 8.72 8.98 7.82 4.82 8.06 6.9 9.7 6.09 4.94 9.27 5.8 9.34 8.8 9.08 5.8 9.12 4.93

46.39 44.93 39.62 36.26 36.41 38.28 43.45 33.35 37.34 36.3 34.14 48.84 34.91 35.98 49.94 46.37 38.69 35 34.35 32.78 31.61 35.97 32.56 36.26 34.12 36.38 31.17 65.92 64.05 57.8 65.66 71.75 63.02 66.89 70.11 61.81 60.18 66.3 68.32 62.64 57.27 67.91 72.24 70.13 57.27 72 73.07

−0.355 −0.256 −0.319 −0.247 −0.252 −0.219 −0.299 −0.322 −0.367 −0.358 −0.206 −0.384 −0.181 −0.173 −0.303 −0.314 −0.188 0.014 −0.293 −0.127 −0.18 −0.324 −0.187 −0.247 −0.09 −0.154 −0.099 −0.155 −0.137 −0.196 −0.269 −0.145 −0.128 −0.223 −0.189 −0.197 −0.217 −0.12 −0.146 −0.196 −0.3 −0.221 −0.158 −0.147 −0.3 −0.133 −0.362

The chromosomal distribution analysis revealed the location of 32 out of 47 BjChis genes on 11 chromosomes of B. juncea, while the remaining 15 genes were found on unanchored scaffolds (S4a). Similarly, in case of C. sativa 70 out of 79 identified CsChis genes were located in 18 chromosomes and the rest nine genes were located on unanchored scaffolds (S4b). These genes are randomly distributed across the genomes of both species, and were detected on 11 out of 18 chromosomes in B. juncea and 18 out of 20 chromosomes in C. sativa. Hence, our results indicated that chitinase genes of both B. juncea and C. sativa are unevenly distributed among the chromosomes of both species.

3.2. Phylogenetic analysis and chromosomal distribution of chitinase genes in B. juncea and C. sativa To further understand the relationship of B. juncea and C. sativa chitinases an unrooted maximum likelihood phylogenetic tree was generated using the protein sequences of both B. juncea and C. sativa plants respectively (Fig. 1). Similarly, the phylogenetic trees were generated using the protein sequences of both plants with known chitinases from A. thaliana (S1a & S1b). Phylogenetic analysis of both species revealed four distinct clusters representing different classes of chitinases (Fig. 1; S1a & b). For example, all chitinases of class-I & classII are grouped in one cluster and that of class-IV, class-V and class-III are classified in separate clusters respectively. However, from the broader viewpoint the five chitinase classes are actually categorized into two mega-groups. The mega-group 1 include all chitinase of GH19 family (class I, II & IV), whereas chitinase of GH18 family (class III & IV) belong to mega-group 2. The chitinases of B. juncea and C. sativa were named on the basis of their known ortholog of A. thaliana, which showed 10 class-I, 6 class-II, 2 class-III, 25 class-IV and 4 class-V chitinases in B. juncea, and 4 class-I, 21 class-II, 3 class-III, 26 class-IV and 25 class-V in C. sativa, respectively.

3.3. Structural analysis of chitinase genes in B. juncea and C. sativa To further understand the structural evolution of the chitinase genes, the BjChis and CsChis genes were subjected to exon-intron organization by comparing the corresponding genomic sequences with their coding sequences (CDS) (Fig. 2a & b). The results revealed that most genes share same number of exons or introns belonging to the same class based on the exon-intron organizations. For example, 13 out of 14 BjChis genes of class-IV had one intron only, and the remaining one has two introns. Similarly, all the BjChis genes of class-V possess 4

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Table 2 Nomenclature and physico-chemical properties of chitinase genes identified in C. sativa. S.No.

Locus

Group

Glyco-hydro Group

Protein length (a.a)

M.wt. (Da)

PI

Instability Index

GRAVY

1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60 61 62 63 64 65 66 67 68 69 70 71 72 73 74

Csa01g013580.1 Csa15g016370.1 Csa15g016380.1 Csa15g016420.1 Csa01g019630.1 Csa02g004830.1 Csa03g002380.1 Csa08g052810.1 Csa13g055600.1 Csa14g002410.1 Csa14g007230.1 Csa15g022910.1 Csa17g001140.1 Csa17g009260.1 Csa19g023850.1 Csa02g068400.1 Csa03g012020.1 Csa10g012890.1 Csa10g016750.1 Csa11g013890.1 Csa11g018230.1 Csa12g017640.1 Csa12g025910.2 Csa14g010070.1 Csa17g012030.1 Csa08g018380.1 Csa13g028570.1 Csa20g039290.1 Csa04g040490.1 Csa04g061310.1 Csa04g061330.1 Csa04g061340.1 Csa04g061350.1 Csa04g061360.1 Csa04g061370.1 Csa04g061380.1 Csa05g006740.1 Csa05g006750.1 Csa05g006770.1 Csa05g006780.1 Csa05g006790.1 Csa05g006820.1 Csa06g028880.1 Csa06g049840.1 Csa06g049870.1 Csa06g049880.1 Csa06g049890.1 Csa06g049900.1 Csa06g049910.1 Csa06g049920.1 Csa09g064130.1 Csa04g030480.1 Csa06g049860.1 Csa09g047920.1 Csa00551s010.1 Csa00551s030.1 Csa00551s040.1 Csa00551s050.1 Csa00656s010.1 Csa02g026860.1 Csa08g034790.1 Csa10g023410.1 Csa10g023410.2 Csa10g023430.1 Csa10g023440.1 Csa10g024440.1 Csa10g024460.1 Csa11g023800.1 Csa11g026520.1 Csa11g026540.1 Csa11g027540.1 Csa11g027550.1 Csa11g027560.1 Csa11g027580.1

I I I I II II II II II II II II II II II II II II II II II II II II II III III III IV IV IV IV IV IV IV IV IV IV IV IV IV IV IV IV IV IV IV IV IV IV IV IV IV IV V V V V V V V V V V V V V V V V V V V V

Glyco_hydro_19 Glyco_hydro_19 Glyco_hydro_19 Glyco_hydro_19 chitinase_glyco_hydro_19 chitinase_glyco_hydro_19 chitinase_glyco_hydro_19 chitinase_glyco_hydro_19 chitinase_glyco_hydro_19 chitinase_glyco_hydro_19 chitinase_glyco_hydro_19 chitinase_glyco_hydro_19 chitinase_glyco_hydro_19 chitinase_glyco_hydro_19 chitinase_glyco_hydro_19 chitinase_glyco_hydro_19 chitinase_glyco_hydro_19 chitinase_glyco_hydro_19 chitinase_glyco_hydro_19 chitinase_glyco_hydro_19 chitinase_glyco_hydro_19 chitinase_glyco_hydro_19 chitinase_glyco_hydro_19 chitinase_glyco_hydro_19 chitinase_glyco_hydro_19 Glyco_hydro_18 Glyco_hydro_18 Glyco_hydro_18 chitinase_glyco_hydro_19 chitinase_glyco_hydro_19 chitinase_glyco_hydro_19 chitinase_glyco_hydro_19 chitinase_glyco_hydro_19 chitinase_glyco_hydro_19 chitinase_glyco_hydro_19 chitinase_glyco_hydro_19 chitinase_glyco_hydro_19 chitinase_glyco_hydro_19 chitinase_glyco_hydro_19 chitinase_glyco_hydro_19 chitinase_glyco_hydro_19 chitinase_glyco_hydro_19 chitinase_glyco_hydro_19 chitinase_glyco_hydro_19 chitinase_glyco_hydro_19 chitinase_glyco_hydro_19 chitinase_glyco_hydro_19 chitinase_glyco_hydro_19 chitinase_glyco_hydro_19 chitinase_glyco_hydro_19 chitinase_glyco_hydro_19 chitinase_glyco_hydro_19 chitinase_glyco_hydro_19 chitinase_glyco_hydro_19 GH18_plant_chitinase_class_V GH18_plant_chitinase_class_V GH18_plant_chitinase_class_V GH18_plant_chitinase_class_V GH18_plant_chitinase_class_V GH18_plant_chitinase_class_V GH18_plant_chitinase_class_V GH18_plant_chitinase_class_V GH18_plant_chitinase_class_V GH18_plant_chitinase_class_V GH18_plant_chitinase_class_V GH18_plant_chitinase_class_V GH18_plant_chitinase_class_V GH18_plant_chitinase_class_V GH18_plant_chitinase_class_V GH18_plant_chitinase_class_V GH18_plant_chitinase_class_V GH18_plant_chitinase_class_V GH18_plant_chitinase_class_V GH18_plant_chitinase_class_V

322 319 331 329 335 280 164 267 280 144 321 337 271 321 335 257 277 235 210 235 210 235 196 280 278 302 302 302 273 282 264 286 263 265 276 279 284 279 266 265 286 278 302 278 286 294 265 281 279 284 273 275 207 275 336 353 358 365 391 348 300 385 379 371 363 368 361 319 379 366 334 363 363 351

34,835.25 35,414.84 35,634.19 35,782.28 36,973.83 31,688.94 17,998.31 30,251.49 31,702.97 15,598.46 35,601.46 37,184.92 30,017.78 35,611.54 36,969.72 28,495.3 30,282.65 25,884.78 23,668.05 25,813.67 23,546.83 25,994.89 21,985.27 30,465.99 30,414.86 33,190.84 33,142.75 33,043.76 29,637.05 30,302.95 28,940.64 31,041.35 28,968.6 28,580.15 30,950.63 29,826.49 30,524.84 29,887.49 28,597.14 29,119 30,724.95 29,845.46 33,190.84 29,827.49 30,787.23 32,219.68 28,539.05 31,741.57 29,877.49 30,566.89 29,532.89 30,626.69 22,448.46 30,456.58 36,991.45 39,284.67 39,792.4 40,582.28 43,631.85 38,675.04 33,714.94 41,845.82 41,167.02 40,090.74 40,183.73 40,686.55 40,504.17 35,412.71 41,153.94 40,148.62 37,332.67 40,011.84 40,263.12 39,432.8

7.81 5.14 7.36 7.81 6.15 8.84 6.81 9.13 8.72 5.56 6.52 5.69 6.42 6.52 5.8 8.77 9.17 9.35 8.95 9 9.06 9.26 9.35 9.25 9.27 9.16 9.08 9.12 4.94 6.17 6.19 4.8 8.44 8.28 8.82 9.66 8.91 9.66 8.28 8.58 4.8 6.7 9.16 6.7 5.1 8.77 7.82 8.98 9.62 8.8 5.18 9.32 6.39 9.33 5.09 4.68 5.29 5.3 5.59 4.75 6.05 8.88 9.07 4.85 5.3 5.22 4.98 4.93 9.08 4.93 4.99 5.01 4.76 4.94

62.24 63.64 62.9 60.91 67.28 67.61 73.17 69.81 69.36 69.17 70.84 68.31 68.82 72.06 68.72 67.16 69.35 82.55 79.38 84.64 79.38 84.21 82.09 74.14 72.95 81.06 82.35 79.14 67.58 34.35 70.98 57.06 63.42 62.98 70 60.61 60.56 57.1 63.12 68.11 56.36 57.01 81.06 58.78 59.44 69.69 63.36 67.05 58.85 59.19 66.15 57.53 70.72 57.53 78.12 73.8 72.29 71.18 72.43 71.72 77.37 76.36 75.25 76.87 71.49 72.8 72.19 71.85 74.72 75.98 74.49 76.34 76.64 69.52

−0.305 −0.384 −0.285 −0.367 −0.281 −0.404 −0.185 −0.357 −0.381 −0.288 −0.231 −0.264 −0.221 −0.202 −0.25 −0.441 −0.331 −0.198 −0.256 −0.146 −0.283 −0.19 −0.245 −0.245 −0.29 −0.163 −0.16 −0.137 −0.135 −0.262 −0.126 −0.381 −0.257 −0.169 −0.234 −0.174 −0.127 −0.214 −0.158 −0.152 −0.343 −0.234 −0.163 −0.206 −0.324 −0.14 −0.158 −0.258 −0.187 −0.152 −0.152 −0.485 0.049 −0.444 −0.194 −0.259 −0.328 −0.341 −0.267 −0.293 −0.375 −0.119 −0.14 −0.089 −0.335 −0.227 −0.378 −0.249 −0.146 −0.276 −0.302 −0.173 −0.244 −0.417

(continued on next page) 5

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Table 2 (continued) S.No.

Locus

Group

Glyco-hydro Group

Protein length (a.a)

M.wt. (Da)

PI

Instability Index

GRAVY

75 76 77 78 79

Csa12g040730.1 Csa12g040750.1 Csa12g040760.1 Csa12g041760.1 Csa13g047080.1

V V V V V

GH18_plant_chitinase_class_V GH18_plant_chitinase_class_V GH18_plant_chitinase_class_V GH18_plant_chitinase_class_V GH18_plant_chitinase_class_V

379 372 365 363 377

41,152.09 40,105.82 40,338.85 40,307.22 41,792.32

9.08 5.44 5.08 4.76 4.99

76.28 75.11 74.05 77.71 69.6

−0.094 −0.108 −0.296 −0.265 −0.374

upstream sequences from both BjChis and CsChis genes revealed many cis-regulatory elements related to biotic stress such as TC-rich repeats (ATTTTC), CGTCA-motif and TGACG-motif (JA responsive element), TCA-element (SA-responsive element), and Box-W1 and EIRE (fungal elicitor responsive elements). Furthermore, many potential elements responsive to biotic stress have been identified such as HSE motif ( CNNGAANNTTCNNG) involved in heat stress, ABREs motif (ACGT) for ABA-dependent expression, MBS/MYB motif (TAACTG) for drought stress, LTRE motif (TGG/ACC GAC) involved in low-temperature response and WUN-motif involved in wound response. Similarly, many hormonal-responsive elements such as ERA (ethylene-responsive element), GARE-motif and P-box (gibberellin-responsive element), TGAelement (auxin-responsive element) found in single or multiple copies in promoters of both BjChis and CsChis genes (S5; Table S3 & S4). In the

two introns, and most BjChis genes of class-I possess one intron. The majority of BjChis genes of class-II and class-III had two introns. Similarly, all the CsChis genes of class-I and class-III had one and two introns respectively, while majority of class-II genes have three introns. The 21 out of 26 class-IV CsChis genes had one intron, and most genes in classV had two exons. Hence, based on exon-intron structure analysis it is reported that chitinases of both crop plants have varied number of exons and introns, and this difference led to variation in gene length as well as their physico-chemical properties (Tables 1 & 2). To determine the regulatory aspect of chitinase genes in response to environmental stresses and hormonal, we analyzed the upstream sequences (1.5 kb) of randomly selected five chitinase genes (one from each class I, II, III, IV and V) from both B. juncea and C. sativa to identify the cis-elements involved in multiple stresses (S5). In silico analysis of

Fig. 1. Phylogenetic relationship among the B. juncea and C. sativa chitinase genes based on the amino acid sequence alignment. The phylogenetic tree is Maximum likelihood (ML) and bootstrap support is based on 1000 replicates. 6

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Fig. 2. a: Analysis of exon-intron structure of B. juncea chitinase genes. Graphic representation of the gene models of 28 BjChis genes identified from B. juncea genome revealed presence of varied number of introns. Exons are shown as green boxes and introns are shown as black lines. b: Analysis of exon-intron structure of C. sativa chitinase genes. Graphic representation of the gene models of 79 CsChis genes identified from C. sativa genome revealed presence of varied number of introns. Exons are shown as blue boxes and introns are shown as black lines. (For interpretation of the references to colour in this figure legend, the reader is referred to the web version of this article.)

basis for understanding the molecular function of chitinase genes in B. juncea and C. sativa. All the chitinases except class-V member of C. sativa have signal peptide at N-terminal with varied length of amino acids, whereas pore linings are present in all the chitinases of both crop species. The signal peptide at the N-terminal assists to deliver the protein at its destination site, and is cleaved off once it reaches its location. Moreover, chitinases of all studied classes are cytoplasmic and their action is extra-cellular (Fig. 3a & b).

upstream regions of chitinase genes presence of these cis-regulatory elements revealed that these genes might be regulated by multiple stresses. In addition, more number of cis-regulatory elements related to defense were present in upstream region of C. sativa chitinase compared to B. juncea (Table S3 & S4).

3.4. Molecular modeling of chitinase genes in B. juncea and C. sativa Molecular modeling of chitinase genes led to insights into the chitin binding domain by providing dynamic and energetic information which usually is difficult to get experimentally. The three-dimensional (3D) protein models were constructed for class I-V chitinases using phyre2 server for understanding the structural properties of chitinase genes in B. juncea and C. sativa, and the results are shown in (Fig. 3a & b). 3D protein models have been constructed with > 90% confidence, and residue coverage varied from 72 to 98. Furthermore, these predicted 3D protein structures are considered highly reliable and offer a preliminary

3.5. Synteny and gene duplication analysis of chitinase gene family The B. oleracea, B. rapa and C. sativa genomes are presently grouped into three sub-genomes, namely LF (least fractionated), MF-I (most fractionated), and MF-II [33]. Our study reported that the C. sativa LF sub-genome possess the most number of chitinase genes (23), followed by sub-genome MF-I (22) and MF-II (17) (Table S5). Each A. thaliana chitinase gene was expected to have three similar copies in Brassica due 7

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Fig. 2. (continued)

orthologous genes of AT1G56680 were not found in C. sativa, and similarly the AT3G47540, AT4G19720, AT4G19730, AT4G19740, AT4G19750, AT4G19760, AT4G19770 genes are lost in B. rapa and B. oleracea. During evolution there have been the expansion in different gene families which occurs through segmental duplication, tandem duplication, and whole-genome duplication [35]. We explored gene duplication events to simplify the genome expansion mechanism of the C. sativa chitinase gene family. The physical distance evaluation between C. sativa chitinase gene loci showed that 12 genes (i.e., Csa04g061310/ Csa04g061400, Csa06g049840/Csa06g049930, Csa05g006740/ Csa05g006730, Csa11g027580/Csa11g026520, Csa10g024460/ Csa10g023410 and Csa12g040730/ Csa12g040730) were tandemly

to Brassica-lineage specific whole-genome triplication (WGT) [34]. However, there were 24 A. thaliana chitinase genes, 36 B. napus chitinase genes and 34 B. oleracea chitinase genes (Table S5). For determining the chitinase genes that have been retained or lost after a WGT event, the syntenic map of C. sativa chitinase genes with the model A. thaliana, B. rapa and B. oleracea chitinase genes provided indicators for defining the sections of conserved synteny among the four genomes (Fig. 4). Equated with the ancestral Brassicaceae blocks (A to X) in A. thaliana, the synteny of 23, 17 and 16 A. thaliana chitinase genes was preserved in C. sativa, B. rapa and B. oleracea, respectively based on the number of corresponding genes (Fig. 4; Table S5). Hence, chitinase family genes were reported to be conserved based on synteny map, along with the duplication or loss of some genes (Fig. 4). The 8

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Fig. 3. a: Predicted 3D structures and transmembrane helix (TM) of five randomly selected B. juncea chitinase proteins, one from each class I-V from top to bottom. 3 b: Predicted 3D structures and transmembrane helix (TM) of five randomly selected C. sativa chitinase proteins, one from each class I-V from top to bottom.

Scavenging enzymes to prevent cell from oxidative burst [77]. At the time of infection by necrotrophic fungal pathogens plants activate different enzymes to counteract the incoming pathogen such as SOD, CAT, POX and glutathione reductase are the most important in neutralizing the effect of superoxide (O2−), or its dismutation product hydrogen peroxide (H2O2). [78]. Superoxide dismutase activity was comparatively higher in C. sativa (30.87 folds) at 12 h but in B. juncea it showed higher percentage at 48 h hpi (Fig. 5) Peroxidase activity was examined in A. brassicae infected leaves of B. juncea and C. sativa at 12, 24 and 48 hpi. The activity of peroxidase was low in B. juncea as compared to C. sativa. Maximum activity of peroxidase was observed at 12 hpi in C. sativa were in B. juncea it showed very little activity in early hour of infection (Fig. 5). C. sativa showed little increase in the peroxidase activity at 12 hpi followed by decline at 24 and 48 hpi. Peroxidase activity decreased successively at 12hpi but increases at 48hpi in B. juncea. Catalase activity was measured from Alternaria infected leaves of B. juncea and C. sativa after 12, 24 and 48 hpi. Catalase acts upon H2O2 produced in the cell during various biotic and abiotic stresses and

duplicated (Table S6). These genes are located on Chr4, Chr5, Chr6, Chr10, Chr11 and Chr12. The data recovered from the BRAD Database showed 19 chitinase genes were associated with segmental duplication that are distributed across the C. sativa genome (Table S6). 3.6. DAB staining and oxidative burst Production of reactive oxygen species is one of the important aspects in plants to respond multiple stresses here we studied the production of H2O2 after the necrotrophic fungal infection (A. brassicae). Microscopic study reveals the production of H2O2 at the infection site due to oxidative burst and it was observed by brown colour. The H2O2 production was more in C. sativa at early hour's (12h) and in B. juncea it increased at late hours. 3.7. Antioxidant enzyme activities Plants have a mechanism of maintaining ROS-homeostasis by ROS 9

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Fig. 3. (continued)

breaks it into water and oxygen. C. sativa showed increased catalase activity at 12 hpi whereas B. juncea the maximum activity (7.40 folds) was at 24 h hpi. Glutathione reductase activity in B. juncea was found to increase at 24 h and then decreased up to 48 h and in C. sativa the activity was maximum at 12 h for pathogen infected plant samples (7.42 fold) and it shows decline at 48 h hpi. (Fig. 5).

Furthermore, the gene expression of C. sativa chitinases showed greater fold change compared to chitinases of B. juncea, which further provides evidence of increased resistance of C. sativa to A. brassicae diseases compared to other Brassica species.

3.8. Transcriptional analysis of chitinase genes in response to Alternaria infection

Chitinases are one of the most important class of PR proteins, and are reported in different living organisms including plants, animals, fungi, and bacteria [37]. They play an essential role in the regulation of growth and development in plants besides being involved in plant defense against fungal pathogens [24]. Being large gene family, it is prerequisite to unravel the function of chitinase genes and to arbitrate their functional specificity for exploitation in genetic analysis and breeding. Although, chitinase gene family have been characterized in many plant species but limited information of chitinase genes are available in B. juncea and C. sativa, and their systematic investigation has not been undertaken. Recent advances in the whole genome sequencing of crop plants have explored the field of functional genomics, which has highly contributed to the identification of stress related genes and their use in genetic engineering as well as molecular breeding. Hence, availability of B. juncea and C. sativa genomic data provides a unique opportunity for identifying putative chitinase genes in these crop species [38,39]. Based on the sequences as well as conserved

4. Discussion

The multiple studies have clearly demonstrated the important role of chitinase genes in antifungal disease resistance in crop plants. In addition, these genes have also been reported to help in growth and development of plants [36]. Therefore, in the present study, we examine the transcriptional changes of chitinase genes of both B. juncea and C. sativa in response to A. brassicae infection (Fig. 6). However, due to the abundance as well as predominant role of class-IV and V chitinases for disease resistance in the family Brassicaceae, we examine the transcriptional changes of six selected chitinase genes of group IV and V (three from each group) from B. juncea (BjChisIV & BjChisV) and C. sativa (CsChisIV & CsChisV). The results of qRT-PCR analysis revealed transcript levels of all the BjChis and CsChis genes increases significantly at 6 h and reached maximum at 24 h of post-inoculation relative to control but their levels decreased sharply at later stages (48 h) (Fig. 6). 10

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Fig. 4. Syntenic relationships of A. thaliana chitinase genes with the chitinases of C. sativa, B. oleracea and B. rapa chromosomes are indicated in blue, orange, violet and green colour respectively. Synteny relationships were lined by Circos (http://circos.ca/). (For interpretation of the references to colour in this figure legend, the reader is referred to the web version of this article.)

Phylogenetic analysis of both B. juncea and C. sativa chitinase sequences separately with that of known chitinases of Arabidopsis clustered them into four well-supported clusters. It is observed that in both cases all the chitinase genes of class-I and class-II of GH 19 are clustered in the same group. This is mainly because GH19 family chitinases adopt the single displacement catalytic mechanism due to high percentage of alpha-helices, whereas all the GH18 family chitinases have triosephosphateisomerise (TIM barrel) as catalytic domains fold with a conserved DxDxE motif, and through substrate-assisted mechanism they catalyze the hydrolytic reaction [46–48]. The class-II was observed to be imbedded within class-I chitinases, because it was suggested that through insertion of chitin-binding domain class-I chitinase may arise from class-II chitinase [49]. Furthermore, the chitinase genes of class III, IV and V are clustered into three separate groups respectively in the phylogenetic trees (S1a & S1b). However, from the broader side the phylogenetic trees of both crop species revealed that all the five classes of chitinase are actually categorized into two mega-groups, which have relatively independent evolutionary history as well as are very distinct to each other (Fig. 1: S1a & b). On the basis of their domain these megagroups can be easily distinguished. The “1” mega-group comprised of classes I, II and IV are characterized by glyco_hydro_19 domain, whereas “2” mega-group possess glyco_hydro_18 domain and include classes III and V chitinases. The GH19 family genes are almost exclusively present in plants, whereas GH18 family genes are distributed in various organisms, including animals, plants, fungi and bacteria [45]. Furthermore, in the present study we identified majority of the

domains, we identified 47 and 79 chitinase genes in B. juncea and C. sativa genome, respectively which is comparatively much higher than reported in A. thaliana [24]. However, the presence of more number of chitinases in B. juncea and C. sativa compared to A. thaliana is attributed to whole-genome triplication (WGT) event that occurred about 13–17 million years ago after the divergence from Arabidopsis [40,41]. Moreover, the number of chitinase genes identified in both crop species are comparatively higher than previously reported in B. rapa (33) [42] and rice (37) [43], but lower than Gossypium hirsutum (92) and Gossypium barbadense (116) [24]. However, B. juncea has almost same number of chitinases as that of Gossypium raimondii (47) and Gossypium arboreum (49) [24], whereas C. sativa has higher number of chitinase genes than E. grandis (67) [44]. Furthermore, it is noted that more number of chitinase genes were found in C. sativa compared to B. juncea, which may be attributed due to the higher ploidy level as well as large genome size of C. sativa [39]. Moreover, chitinase genes in both B. juncea and C. sativa are unevenly distributed present in 11 of the 18 and 18 of the 20 chromosomes, respectively (Fig. S3a & b). These findings are similar to that reported in B. rapa were 33 chitinase genes are present on 8 out of total 10 chromosomes[42]. Jiang et al. [45]also revealed that 37 chitinase genes in P. trichocarpa are located on eight of the 19 chromosomes. Similarly, in rice 37 chitinase genes have been reported to be distributed unevenly [43]. In our lab we have validated the role of chitinase class-IV gene by overexpressing in B. juncea transgenic plants in response to A. brassicae, which showed the improved resistance against this pathogen (data not shown). 11

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Fig. 5. SOD, CAT, POX and Glutathione Reductase activity in B. juncea and C. sativa at 12, 24 and 48 hpi after A. brassicae infection. Microscopic analysis of DAB Stained B. juncea and C. sativa leaves at 0, 12, 24 and 48hpi.

which involves presence of single or multiple copies of ABREs motifs, or independently that possess DRE motifs (TAC CGA CAT) binding to different DREBPs groups [56]. The presence of drought-inducibility elements/motifs viz., MYB and MBS in the studied upstream regions implying their regulatory role in drought response [57]. In addition, LTRE commonly present in cold-responsive genes is a low-temperatureresponsive element motif [58], whereas in the promoters of heat shock protein genes (HSPs) are the cis-acting sequences designated as heat shock elements (HSEs) that led to rapid expression of HSP, which in turn result enhanced thermo-tolerance [59]. The presence of hormone responsive elements viz., ERA (ethylene-responsive element), GAREmotif and P-box (gibberellin-responsive element), TGA-element (auxinresponsive element) in chitinase gene promoter's revealed their important role in regulation of chitinase genes. In a wide range of biotic and abiotic stresses presence of SA- and JA-responsive motifs in stressrelated genes are reported to enhance stress tolerance [60]. The B. juncea and C. sativa PR genes are known to possess these motifs in single or multiple copies, and thus further confirmed about their important role in multiple stress [60]. However, the more number of defense-related cis-regulatory elements in upstream regions of C. sativa chitinase genes than B. juncea provides further evidence about their potential role in disease resistance. Since after speciation from A. thaliana, the genome of the members of family Brassicaceae has undergone whole-genome triplication (WGT) [61,62], the C. sativa genome theoretically contains three copies of A.

GH 19 family chitinases as compared to those belonging to GH 18 family in both crop species, which is similar to that reported in other crop species viz., B. rapa [42], Musa acuminata [50] and Zea mays [51] etc. In general, fewer introns are possessed by stress-related genes [52], similarly in the present study 27 of the 28 and 52 of the 79 chitinase genes identified in B. juncea and C. sativa, respectively has two or fewer introns confirmed this conception (Fig. 2a & b). Other stress-related gene families such as leucine-rich repeat (LRR) family [53], LEA family [54], and the trehalose-6-phosphate synthase gene family [55] also contain low intron numbers. Moreover, transcription is prolonged due to long and large number of introns, whereas genes possess only few introns are regulated rapidly during stress [52]. Hence, less number of introns in chitinase genes will allow their quick transcription as well as action following the pathogen attack. To further elucidate the expression of chitinase genes under stress in B. juncea and C. sativa, 1.5 kb upstream regions were used for scanning of cis-elements involved in multiple stresses in both crop species. Based on in silico analysis, many cis-acting regulatory elements related to stress present in single or multiple copies were observed in the upstream sequences. Among the cis-regulatory elements related to biotic stress are TC-rich repeats, JA (TGACG & CGTCA) motifs, SA (TCA) motifs, and fungal elicitor responsive elements (Box-W1 and EIRE) present in gene promoters of chitinases, and thus confirmed the important role of chitinase genes in biotic stress. Genes related to abiotic stress are usually activated either through ABA-dependent pathway

12

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Fig. 6. Expression analysis of randomly selected six chitinase genes (three from each class-IV and -V) at 6, 12, 24 and 48 h after A. brassicae infection in both B. juncea and C. sativa. Three biological replicates were used to calculate error bars, based on standard error (*P < .05; **P < .01)

thereby allows cells to resist peroxidation of membrane lipids and maintain integrity of the cell membrane [80]. In the present study, A. brassicae infected B. juncea and C. sativa plants increased the activity of SOD, CAT, POX and GR which in turn reduced the overproduction of ROS to reduce oxidative stress (Fig. 5). SOD activity was more in C. sativa than B. juncea. It was reported to be higher at 12 h hpi in C. sativa than B. juncea. There are only few studies on enzyme activities after necrotrophic fungal infection in plants and here we demonstrated first time the possible role of SOD, CAT, POX and GR after necrotrophic fungal infection by A. brassicae in B. juncea and C. sativa our results were consistent with [81] were they showed the enzyme activities against A. alternata. Different PR proteins have been revealed to play a key role in plant immune system by modulating plant defense system. In this regard, the chitinases are well known as PR proteins belonging mostly to PR-3 family [2]. The previous studies have revealed that both biotic and abiotic stresses activate the expression of PR3 [24,66]. Therefore, in the present study our results revealed that the expression of class-IV and class-V chitinases was significantly induced by A. brassicae infection and reached peak at 24 h after inoculation in both B. juncea and C. sativa. Previous studies also revealed a similar expression pattern of chitinase genes, however there were differences in transcript levels, which might be due to the type of plant-pathogen interactions or by nature [6,67]. Expression profile of PR3 gene in A. thaliana and B. napus inoculated with A. brassicicola and Phoma. lingarn, respectively also support our findings [68,69]. Similarly, Xu et al. [24] reported that chitinase genes are significantly induced after Verticillium dahlia inoculation in cotton and expression level of these genes reached the

thaliana chitinase genes. However, all the A. thaliana chitinase genes have their homologous in the C. sativa genome except AT1G56680 chitinase gene. Moreover, A. thaliana chitinase genes viz., AT3G47540, AT4G19720, AT4G19730, AT4G19740, AT4G19750, AT4G19760 and AT4G19770 showed duplication in C. sativa, whereas rest of the 16 A. thaliana chitinase genes revealed triplication. This result provides evidence that before speciation these genes were conserved, but due to artificial selection or evolution these genes might have lost. In other Brassica species such loss of chitinase genes was also observed, for example, Chen et al. [42] reported that either six genes were not duplicated or 10 genes orthologous to Arabidopsis chitinases were lost in B. rapa. Similarly, losses of genes during the WGT event have been found in MKK and MPK genes (Pioa et al. 2018), nucleotide binding site (NBS)-encoding genes [64], late embryogenesis-abundant (LEA) genes [54], isopentenyl transferase (IPT) and cytokinin oxidase/dehydrogenase (CKX) genes [65] of Brassica species. Thus, in C. sativa genome loss or expansion of some chitinase family genes indicates that under different environmental stress conditions these genes has undergone some functional differentiation. During the long evolutionary process, the C. sativa has possibly sufficient number of chitinase genes to resist outer stresses. The SOD, CAT, POX, and Glutathione reductase are the key enzymes which detoxify the ROS at the time of infection [79]. However, SOD is believed to serve as a frontline antioxidant defense against biotic and abiotic stresses by detoxifying O2 that results formation of H2O2 (damaging cell organelles, protein and nucleic acids), that in turn is detoxified by CAT and POX. The POX has significant importance in reducing H2O2 accumulation, eliminating MDA (malondialdehyde) 13

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highest peak at 24 hpi, suggesting their effective role in disease resistance. The present study also revealed greater fold change in expression of C. sativa chitinase genes compared to B. juncea after pathogen inoculation, which further confirms the enhanced resistance of C. sativa to A. brassica. These findings are supported by previous studies where C. sativa have been reported to show higher resistance against A. brassicae compared to other Brassica species [21,22,70].

[3] [4]

[5] [6]

5. Conclusions [7]

In conclusion, this is the first comprehensive study and systematic genome-wide analysis of chitinase gene families in B. juncea and C. sativa. Hence, we identified 47 and 79 chitinase genes in B. juncea and C. sativa, respectively which were unevenly distributed on chromosomes in both crop species. Based on amino acid sequence similarities, chitinases of both species were classified into five classes (I-V), and these classes are clustered into four major groups of phylogenetic trees. Members of the same group or subgroup reveal highly conserved exon/ intron composition, conserved domain, and motifs. Furthermore, the synteny analysis revealed that chitinase gene family in C. sativa, B.rapa and B. olearacea crop plants have expanded through WGT, segmental and tandem duplication events, and this gene family is under positive selection pressure. Enzyme assays also revealed the possible role in defense in response to A. brassicae infection. The qRT-PCR expression analysis revealed that chitinase genes are significantly induced by A. brassicae infection, and thus provides insights about their potential role in disease resistance. Moreover, chitinase genes of C. sativa showed greater fold change in expression compared to B. juncea following A. brassicae inoculation and provides further evidence for increased disease resistance of C. sativa. This fact has been further proved as more number of defense-related cis-elements were present in upstream regions of chitinase genes of C. sativa compared to B. juncea. Hence, this leads to the conclusion that C. sativa chitinase genes can be effectively used for the development of disease resistant varieties of Brassica crop species.

[8] [9] [10] [11] [12] [13]

[15] [19] [20]

[21]

[22]

[23]

[24]

Author contributions

[25]

AG and AS conceived and designed the research. ZAM has performed all the experiments as well as wrote the manuscript. SA, JAB, SMS, SR and PKP contributed to data analysis. JAB, SMS, SA, and PY has contributed in bioinformatic analysis. AG and SA contributed in manuscript proofreading. All authors read and approved the manuscript.

[26]

[27]

[28]

Competing interests [29]

Authors declare no competing interests.

[30] [31]

Acknowledgments

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We gratefully acknowledge the Project Director, National Research Centre on Plant Biotechnology for providing all the facilities and also ICAR-NPTC for financial support to complete this work.

[34]

Appendix A. Supplementary data

[35] [36]

Supplementary data to this article can be found online at https:// doi.org/10.1016/j.ygeno.2019.05.011.

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