Poplar carbohydrate‐active enzymes: whole‐genome annotation and functional analyses based on RNA expression data

Summary

Carbohydrate‐active enzymes (CAZymes) catalyze the formation and modification of glycoproteins, glycolipids, starch, secondary metabolites and cell wall biopolymers. They are key enzymes for the biosynthesis of food and renewable biomass. Woody biomass is particularly important for long‐term carbon storage and as an abundant renewable natural resource for many industrial applications. This study presents a re‐annotation of CAZyme genes in the current Populus trichocarpa genome assembly and in silico functional characterization, based on high‐resolution RNA‐Seq data sets. Altogether, 1914 CAZyme and expansin genes were annotated in 101 families. About 1797 of these genes were found expressed in at least one Populus organ. We identified genes involved in the biosynthesis of different cell wall polymers and their paralogs. Whereas similar families exist in poplar and Arabidopsis thaliana (with the exception of CBM13 found only in poplar), a few families had significantly different copy numbers between the two species. To identify the transcriptional coordination and functional relatedness within the CAZymes and other proteins, we performed co‐expression network analysis of CAZymes in wood‐forming tissues using the AspWood database (http://aspwood.popgenie.org/aspwood-v3.0/) for Populus tremula. This provided an overview of the transcriptional changes in CAZymes during the transition from primary to secondary wall formation, and the clustering of transcripts into potential regulons. Candidate enzymes involved in the biosynthesis of polysaccharides were identified along with many tissue‐specific uncharacterized genes and transcription factors. These collections offer a rich source of targets for the modification of secondary cell wall biosynthesis and other developmental processes in woody plants.

Significance Statement

Re‐annotation of carbohydrate‐active enzymes in poplar and their transcript analyses reveal genes and gene families involved in wood cell wall biosynthesis.

Introduction

Carbohydrate‐active enzymes (CAZymes) are essential proteins for all life forms (Lombard et al., 2014). They are fundamentally important for the synthesis, degradation and modification of all the glycoconjugates found in nature, including carbohydrates, glycoproteins, glycolipids and glycosylated secondary metabolites (Wilson, 2002; Cantarel et al., 2009). Plant tissues are particularly rich in complex carbohydrates, building the cell walls and thus affecting growth and development. Therefore, plants depend heavily on carbohydrate metabolism for a diverse array of physiological processes, including growth, defense, dormancy, signaling and photosynthesis (Coutinho et al., 2003). As a result of high abundance and sustainable carbon assimilation from the atmosphere, the carbohydrate‐rich tissues of plants have become important feedstocks for biofuel and biomaterial production (Himmel et al., 2007).

The classification of CAZymes into families is a difficult task, as different parameters need to be considered simultaneously, including a significant similarity of amino acid sequences with at least one biochemically characterized family member, the presence of defined catalytic and non‐catalytic modules, and the availability of a full‐length protein sequence (Lombard et al., 2014). By adopting this approach, the CAZymes were previously classified into four major classes, namely glycosyltransferases (GTs), glycoside hydrolases (GHs), polysaccharide lyases (PLs) and carbohydrate esterases (CEs) (Cantarel et al., 2009). Recently, the lignin degrading enzymes and the polysaccharide lytic monooxygenases have been added and classified as auxiliary activity (AA) families, which encompass a broad range of enzymes related to lignocellulose conversion (Levasseur et al., 2013). In plants, this group also contains diverse enzymes involved in biosynthesis and developmental processes. In addition, the carbohydrate binding modules (CBMs) encompass a group of proteins with specific domains that bind to carbohydrates (Boraston et al., 2004), for example to substrates, as in the case of hydrolytic enzymes (Guillen et al., 2010), or to ligands, as in in case of receptors (Galindo‐Trigo et al., 2016). These six categories in total are organized in sequence‐based families in the CAZy database (http://www.cazy.org).

As primary producers of terrestrial ecosystems, woody plants play a special role in providing an ecologically essential carbon sink, as well as serving as an important source of industrial raw material. A major proportion of assimilated carbon in woody species is used for cambial growth and the development of the secondary cell walls in xylem cells. Once the carbon is allocated to cellulose, lignin or, to a large extent, hemicelluloses in xylem cell walls, it is immobilized for many decades or even centuries. Therefore, the carbohydrate‐rich secondary cell walls of woody biomass are particularly important as a renewable natural resource for CO₂‐neutral industrial applications, with the goal of replacing fossil‐based resources (Grattapaglia et al., 2009; Hinchee et al., 2009). As CAZymes are involved in the formation and modification of this carbohydrate matrix (Mellerowicz and Sundberg, 2008), a detailed study of CAZymes in a woody model species and their spatial interactions during wood cell wall formation is essential to inform tree improvement programs to support the future bioeconomy.

The availability of a mature genome assembly (v3.0) for Populus trichocarpa, together with recent high‐resolution RNA‐Seq data sets for Populus sp. (Sundell et al., 2015, 2017; Immanen et al., 2016), prompted us to carry out a re‐annotation and expression analysis of CAZymes in Populus. Here, we redefined all members of different CAZyme families and closely related expansins, and compared their genetic diversity with our earlier P. trichocarpa v1.0 annotation (Geisler‐Lee et al., 2006) and with the current Arabidopsis thaliana (http://www.cazy.org) annotation. Furthermore, we have annotated CAZyme families in P. trichocarpa and their orthologs in A. thaliana. The available Populus RNA‐Seq data sets were used to determine tissue specificity, and the co‐expression network analyses among the CAZymes and other genes enabled the discovery of additional gene models with putative function in wood cell wall biosynthesis.

Results and discussion

Identification of CAZyme families

The comparison of protein‐coding transcripts of P. trichocarpa gene models (v3.0) with CAZyme and expansin modules in the CAZy database resulted in the identification of 1914 CAZyme and expansin gene models, including 629 GHs, 788 GTs, 42 PLs, 106 CEs and 65 expansin‐related sequences. Two other CAZyme superfamilies, the recently added catalytically active AAs (112) and the non‐catalytic CBMs (173, excluding those appended to GH and GT families), were also identified. All of the genes identified are listed in Tables S1–S7.

Compared with the previous annotation (Geisler‐Lee et al., 2006), the current annotation includes more families in GTs, GHs and CEs. Thus, we have identified 42 out of 106 GT families, 37 out of 156 GH families, two out of 29 PL families, four out of 16 CE families, four out of 15 AA families and 12 out of 84 known CBM families listed in the CAZy database (as found in November 2018) (Table 1). Populus has all the CAZyme families identified in A. thaliana, whereas one family found in Populus, CBM13, does not have counterparts in A. thaliana (Table 1). This domain has a trefoil structure, with three sugar binding domains that can harbor different specificities to mono‐ or disaccharides and that belong to the ricin B lectin family (Fujimoto, 2013). All identified P. trichocarpa CBM13 gene models clustered at two loci on chromosome 6, and were similar to ricin and abrin‐a, potent plant toxins.

Table 1.

CAZyme families and the number of CAZymes models per family detected in Populus trichocarpa v3.0 and Arabidopsis thaliana v10.0 genomes (November 2018)

CAZyme families	Ath	P. trichocarpa		Ath vs Pt Q values	CAZyme families	Ath	P. trichocarpa		Ath vs Pt Q values
	v.10	v.3	v.3 vs v.1			v.10	v.3	v.3 vs v.1
AA1	36a	78a	78	3.3*	(Continued)
AA5	7a	11a	11	0.1	GH63	2	1	−2	0.9
AA6	4a	9a	9	0.5	GH77	2	3	0	0.1
AA7	27a	14a	14	11.6**	GH79	3	6	−1	0.2
CE6	2	4a	4	0.2	GH81	2	1	−3	0.9
CE8	67	89	0	0.7	GH85	2	2a	2	0.2
CE11	4a	2a	2	1.8	GH89	1	3	1	0.4
CE13	12	11	1	1.5	GH95	1	4	0	0.9
PL1	26	31	3	0.8	GH100	9	16	−3	0.2
PL4	8	11	0	0.1	GH116	4	5a	5	0.1
EXPN	36	65	23	0.8	GH146	2	3a	3	0.1
(CBM13)1–2	0	6	6	4.0**	GH152	19	40a	40	1.5
CBM18‐(EXPN)0–1	1	5	5	0.9	GT1	122	281	−45	15.6**
CBM18‐GH19	9	15a	15	0.1	GT2	42	60	−14	0.1
CBM20	2	7a	7	1.2	GT4	24	38	−3	0.1
(CBM20)2‐GH77	1	2a	2	0.1	GT5	6	7	−6	0.3
(CBM22)1–4‐GH10	11	8a	8	2.6	GT8	42	56	−5	0.4
CBM32	1a	1a	1	0.1	GT10	3	4	0	0.1
CBM43	31	34a	34	1.7	GT13	1	2	−1	0.1
GH17‐CBM43	28	43a	43	0.1	GT14	11	16	2	0.1
CBM45	2	2a	2	0.2	GT16	1	1	0	0.1
(CBM45)2‐GH13	1	1a	1	1.6	GT17	7	5	0	1.8
CBM48	8	18a	18	0.9	GT19	1	1	0	0.1
(CBM48)1–2‐GH13	7	10	10	0.1	GT20	11	13	0	0.4
GH9‐CBM49	3	3a	3	0.3	GT22	3	5	1	0.1
CBM50	1	12a	12	5.7**	GT24	1	1	−1	0.1
(CBM53)3‐GT5	3	1a	1	2.1	GT28	4	5	−1	0.1
CBM57	4	92a	92	51.1**	GT29	3	5	0	0.1
GH1	48	49	−1	3.8*	GT30	1	1	0	0.1
GH2	2	3	0	0.1	GT31	33	48	−4	0.1
GH3	16	27	−6	0.2	GT32	6	5	−1	1
GH5	13	24	2	0.4	GT33	1	1	0	0.1
GH9	29	32	−1	1.6	GT34	8	9	1	0.4
GH10	12	8	1	3.4*	GT35	2	5	−1	0.4
GH13	10	16	−5	0.1	GT37	10	7	−1	2.6
GH14	9	14	2	0.1	GT41	2	4	−1	0.2
GH16	33	43	2	0.5	GT43	4	7	0	0.1
GH17	51	88	−9	0.6	GT47	39	70	−5	0.8
GH18	11	28	−2	2.3	GT48	13	13	−6	1.2
GH19	14	21	0	0.1	GT50	1	1	0	0.1
GH20	3	6	0	0.2	GT57	2	3	−1	0.1
GH27	4	9	1	0.5	GT58	1	1	0	0.1
GH28	68	85	−4	1.4	GT59	1	1	0	0.1
GH29	1	3	0	0.4	GT61	8	10	0	0.2
GH31	5	12	−2	0.8	GT64	3	5	1	0.1
GH32	8	8	−1	0.7	GT66	2	4	−2	0.2
GH33	1	1a	1	0.1	GT75	5	10	−1	0.3
GH35	18	23	1	0.3	GT76	1	2a	2	0.1
GH36	6	17	−2	1.9	GT77	19	19	3	1.7
GH37	1	4	0	0.9	GT90	9	7a	7	1.8
GH38	4	9	1	0.5	GT92	5	3a	3	1.8
GH43	2	1	−1	0.9	GT95	3	6a	6	0.2
GH47	5	6	‐1	0.2	GT96	2	1a	1	0.9
GH51	2	6	3	0.8	GT106	34	44a	44	0.5

^aFamilies in P. trichocarpa not reported by Geisler‐Lee et al. (2006) or in A. thaliana not listed in the CAZy database (http://www.cazy.org/). The identified A. thaliana genes include CBM32 (At5g49570) and AA family genes listed in Table S8.

* and **, families differing in relative gene frequencies between P. trichocarpa and A. thaliana, taking into account the total number of genes in these species, 41377 and 27416, respectively (*P ≤ 0.10; **P ≤ 0.05, χ² test).

Major changes of CAZyme gene families in P. trichocarpa v3.0 versus v1.0 assembly

The current P. trichocarpa v3.0 genome contains more CAZymes and expansin gene models (1914) than were reported for the previous v1.0 assembly (1603; Geisler‐Lee et al., 2006). This is essentially linked to the expansion of the CAZyme database, in which several families have been added or expanded since the first analysis. Considering the previously annotated families (Geisler‐Lee et al., 2006), the total number of genes has been reduced by 7% on average. For example, GT2 lost 14 members, whereas GT8 and GT47 each lost five gene models. This reduction of family members is the result of the significantly improved method of coding sequence detection and improved genome assembly. The lists of v1.0 gene models annotated as CAZymes coinciding (at least partially) with the current CAZyme models are included in Tables S1–S4 and S6, and the change in the number of genes in each family is listed in Table 1.

Genetic diversity of CAZyme gene families in Populus compared with A. thaliana

Approximately 1.6 times more CAZyme genes were identified in P. trichocarpa (v3.0) than in A. thaliana, as listed in the CAZy database in November 2018 (http://www.cazy.org) and supplemented by our homology‐based annotation of A. thaliana AAs (Table S8). This increase was similar to that previously reported (Geisler‐Lee et al., 2006). This observation is in line with the census of CAZyme motifs carried out in the genomes of several streptophytes, including green algae, a moss and many vascular plants, and shows that whereas the woody plants appear to have the highest absolute numbers of CAZyme motifs in their genomes, the frequencies of these domains in relation to respective genome size are stable (Pinard et al., 2015). Thus, the difference in the absolute number of CAZymes between Populus and A. thaliana is related to the linage‐specific history of whole‐genome duplication and gene loss (Tuskan et al., 2006).

Nevertheless, the comparative analysis of relative gene abundance (in relation to genome size) in each CAZyme family between A. thaliana and P.  trichocarpa showed significant differences in a few families (Table 1). For example, the GT1 family, responsible for the glycosylation of different metabolites, the AA1 family, comprising lignification‐related laccases, the CBM50 family, known also as LysM domain binding chitin, and the CBM57 family, including various leucine‐rich receptor‐like kinases (LRKs) were over‐represented in P. trichocarpa. Conversely, the GH1 family of diverse exoglycanases, the GH10 family containing xylanases, the GH17 family involved in callose metabolism and the AA7 family comprising the berberine bridge enzymes were proportionally over‐represented in A. thaliana. These differences provide data for hypotheses on adaptations to perennial/woody versus annual/herbaceous lifestyles, which can be tested by comparative genome analyses.

CAZyme families identified in Populus since the last whole‐genome annotation

Glycosyltransferases

The glycosyltransferases (GTs; EC 2.4.X.X) play a major role in the biosynthesis of cell wall polymers and starch, and the glycosylation of proteins, lipids and secondary metabolites. We annotated six GT families in the P. trichocarpa v3.0 genome assembly that were not annotated in v1.0 assembly (Geisler‐Lee et al., 2006) (Table 1). The GT76 family includes fungal and human mannosyl transferases that use dolichol‐P‐mannose as a donor to synthesize the glycophosphatidylinositol (GPI) anchor (Kang et al., 2005). The GPI anchor proteins are abundant in plants, and GT76 might have a similar role in this kingdom. The GT90 family includes β‐1,2‐xylosyltransferases involved in protein glycosylation and in the biosynthesis of glucuronoxylomannan and galactoxylomannan found in fungal capsules (Klutts et al., 2007).

The remaining four GT families, GT92, GT95, GT96 and GT106, have been created since the previous Populus CAZyme analysis and they represent different activities based on studies in A. thaliana. GT92 members are UDP‐galactose 4‐galactosyltransferases that synthesize rhamnogalacturonan‐I (RG‐I) β‐1,4‐d‐galactan (Liwanag et al., 2012). GT95 members have been characterized as hydroxyproline O‐arabinosyl transferases (HPATs) catalyzing the transfer of l‐arabinofuranosyl (Araf) residue from UDP‐β‐L‐Araf to the hydroxyl group of the hydroxyproline residues of extensins (Ogawa‐Ohnishi et al., 2013), peptide hormones (Schnabel et al., 2011; Okamoto et al., 2013; Xu et al., 2015) and possibly other proteins. The GT96 family comprises serine O‐α‐galactosyltransferases (SGTs) (Saito et al., 2014) involved in the post‐translational modification of arabinogalactan proteins (AGPs) and extensins. It is represented by a single model in P. trichocarpa, and the protein was previously annotated in the CAZy database (http://www.cazy.org). GT106 is a recently created CAZyme family, one clade of which contains RG‐I:rhamnosyltransferases (RRTs) involved in the biosynthesis of the RG‐I backbone (‐4GalUAα1‐2Rhaα1‐)_n (Takenaka et al., 2018). Three out of 44 GT106 Populus sequences had previously been deposited in the CAZy database (http://www.cazy.org).

Glycoside hydrolases

Glycoside hydrolases (EC 3.2.1.X) catalyze the hydrolysis of O‐ or S‐glycosidic bonds and have a major role in cell wall remodeling and architecture, carbohydrate metabolism and protein post‐translational modifications, affecting the physiological activities of plant cells. Five additional GH families were annotated since the previous analysis (Geisler‐Lee et al., 2006) (Table 1). One of these, GH85, has been characterized in A. thaliana as cytosolic endo‐N‐glucosaminidase that cleaves O‐glycosidic linkage between two N‐acetylglucosamines of N‐glycans (Fischl et al., 2011). Four other families have been studied in organisms other than plants. GH33 comprises sialidases (EC 3.2.1.18) cleaving α‐ketosidic linkage between the sialic (N‐acetylneuraminic) acid and a sugar residue in prokaryotes, fungi and animals. Sialic acid is not found in plants, but its derivatives, KDO and 2‐keto‐3‐deoxy‐d‐lyxo‐heptulosaric acid (DHA) are present in the side chain C and D of rhamnogalacturonan‐II (RG‐II), and could perhaps serve as substrates for plant GH33 members. The GH116 family includes diverse activities such as β‐glucosidase (EC 3.2.1.21), β‐xylosidase (EC 3.2.1.37), acid β‐glucosidase/β‐glucosylceramidase (EC 3.2.1.45) and β‐N‐acetylglucosaminidase (EC 3.2.1.52), and is found in all life domains. GH146 and GH152 are recently created GH families in the CAZy database. The GH146 family comprises β‐l‐arabinofuranosidase (EC 3.2.1.185) characterized in bacteria of human gut cleaving both β‐1,2‐ and β‐1,3‐linked l‐Araf branches recently identified in pectic arabinan (Luis et al., 2018). The GH152 family groups stress‐induced thaumatin‐like proteins that are highly abundant in plants (Trudel et al., 1998). Seven out of 40 Populus GH152 members have already been annotated in the CAZy database (http://www.cazy.org). A fungal GH152 member has been shown to have β‐1,3‐glucanase (EC 3.2.1.39) activity (Sakamoto et al., 2006). Plant GH152 proteins exhibit callose‐binding activity (Trudel et al., 1998), suggesting that they could also be β‐1,3‐glucanases.

Carbohydrate esterases

The carbohydrate esterases (CEs) are involved in the remodeling and degradation of the plant cell wall polysaccharides. Plant genomes are known to contain pectin methyl esterases (PMEs, EC 3.1.1.11) in family CE8, and pectin acetyl esterases (EC 3.1.1.X) in family CE13, with both families being previously annotated in P. trichocarpa (Geisler‐Lee et al., 2006). Since then, one member of family CE6 has been annotated in Populus (http://www.cazy.org), whereas we report here three additional CE6 members (Table 1). CE6 contains microbial xylan acetyl esterases (Neumüller et al., 2015), but its function in plants has not been demonstrated. CE11 members are annotated in A. thaliana as UDP‐3‐O‐acyl N‐acetylglucosamine deacetylases that are likely to be involved in lipid biosynthesis via a recently discovered pathway resembling that of lipid A biosynthesis in bacteria (Li et al., 2011a). We have identified two members of CE11 in P. trichocarpa (Table 1). The CAZy database additionally lists one CE4 member annotated as Populus trichocarpa × deltoides ABK96767. This sequence is not homologous to any gene model in P. trichocarpa v3.0, but is instead highly similar to a rust Melampsora larici‐populina 98AG31 CE4 protein XP_007418383. Thus, it appears that the CE4 family is not present in Populus.

Subfamilies of polysaccharide lyases

Polysaccharide lyases (PLs) (EC 4.2.2.‐) cleave 4‐O‐glycosidic bonds in pectins and glucosaminoglycans using a β‐elimination mechanism without using a water molecule (Yip and Withers, 2006). They are involved in diverse biochemical processes, including biomass degradation, cell wall matrix recycling and pathogenesis, and are largely substrate specific (Herron et al., 2000; Abbott and Boraston, 2008). Plants have members in PL1, characterized as pectate lyases (PELs; EC 4.2.2.2) (Domingo et al., 1998; Wang et al., 2010; Biswal et al., 2014), and in PL4, annotated as rhamnogalacturonan lyases (EC 4.2.2.23), that affect cell separation and pectins in middle lamella (Molina‐Hidalgo et al., 2013), but the actual activity has not been demonstrated for any PL4 plant protein. The lack of critical catalytic residues in many of the plant PL4 proteins has led to doubts regarding their enzymatic activity (Kozlova et al., 2017). Based on sequence similarity and substrate specificity, the PL families were further divided into subfamilies represented by an Arabic numeral following the family identifier (Lombard et al., 2010). The current Populus CAZyme annotation identified PL1_1, PL1_12, and PL4_2 subfamilies (Table S3). PL1_1 and PL1_12 both include pectate lyases but PL1_12, classified as clade V of the pectate lyase family, does not have a Ca²⁺ binding site or other motifs present in PL1_1 (Bai et al., 2017).

Auxiliary activities

The category broadly termed as ‘auxiliary activities’ (AAs) has been introduced to the CAZy database since the previous Populus genome‐wide annotation (Geisler‐Lee et al., 2006). It groups together the families of redox enzymes and lytic polysaccharide monooxygenases (LPMOs) involved in polysaccharide, oligosaccharide and lignin breakdown, lignin polymerization and other metabolic processes (Levasseur et al., 2013). The P. trichocarpa v3.0 genome assembly contains four AA families (Tables 1 and S5), similar to A. thaliana. Not all A. thaliana members have been annotated in the current CAZy database, however. Therefore, P. trichocarpa sequences were used to identify the corresponding proteins in A. thaliana by amino acid sequence alignment and phylogenetic analysis (Figure S1; Table S8).

The AA1 family, annotated as multicopper oxidase, is the most abundant Populus AA family (Figure S1; Table 1). It includes a large subfamily of laccases that are considered as key lignin polymerizing enzymes, possessing p‐diphenol:O₂ oxidoreductase activity (EC 1.10.3.2), although only a few members have been shown to carry out this function (Turlapati et al., 2011; Lu et al., 2013), and a large subfamily of SKU5‐SIMILAR (SKS) proteins with GPI anchor that are implicated in stress responses and the regulation of development (Sedbrook et al., 2002). Forty‐nine laccases were annotated in P. trichocarpa (Lu et al., 2013) and our analysis identified 10 additional gene models, some of which might be truncated (Table S5). The AA1 family is abundantly represented in the CAZy database, with 56 entries for P. trichocarpa and several for Populus alba, Populus fremontii, Populus nigra and Populus tomentosa (http://www.cazy.org). Closer inspection of P. trichocarpa entries, however, reveals that they are all partial sequences and constitute multiple copies of four laccase proteins.

The AA5 family, comprising 11 members, contains extraplasmic copper radical oxidases of unknown function, with similarity to glyoxal oxidases, and using oxygen as the acceptor to oxidize aldehydes, with the generation of hydrogen peroxide (Daou and Faulds, 2017). The plant members studied so far were implicated in defense (Guan et al., 2010), seed coat mucilage cohesion and cell adhesion (Šola et al., 2019), and pollen development (Phan et al., 2011). The proposed enzymatic activity of one characterized member, RUBY, was the RG‐I galactose oxidase (Šola et al., 2019). One of the 11 identified AA5 members has been previously annotated in P. tomentosa as glyoxal oxidase in the CAZy database (http://www.cazy.org).

The AA7 family, containing 14 members in P. trichocarpa, comprises glucooligosaccharide oxidases (GOOs) that oxidize the anomeric carbon hydroxyl groups of glucose or α‐ and β‐1,4‐linked sugars, first to lactone and then to the corresponding acid, and reduce O₂ to H₂O₂ (Levasseur et al., 2013). It includes plant apoplastic proteins named as berberine bridge enzymes and nectarins, which are thought to function in defense responses (van Hellemond et al., 2006; Benedetti et al., 2018). They have been implicated in the oxidation of oligogalacturonides, functioning as damage‐associated molecular patterns (DAMPs) (Benedetti et al., 2018), and possibly in lignin polymerization by the oxidation of monolignols (Daniel et al., 2015).

The AA6 family, with six members in P. trichocarpa, contains intracellular 1,4‐benzoquinone reductases, which might be involved in detoxification. One member in A. thaliana, AtFRQ1, was found to be rapidly induced by auxin (Laskowski et al., 2002).

Carbohydrate binding modules

Carbohydrate binding modules (CBMs) are non‐catalytic domains with autonomous folding that recognize specific carbohydrate motifs (Boraston et al., 2004). They can be part of enzymes and receptors or can make an entire protein like OLE E 10 (CBM43), found in the pollen of the olive tree (Barral et al., 2005). Among all CBMs, CBM57 was the most abundant in Populus (Table 1). This domain, known as the malectin domain in animals, has been implicated in a protein glycosylation surveillance mechanism in the endoplasmic reticulum (ER) (Schallus et al., 2008). In plants, CBM57 is found in receptor‐like kinases (RLKs) (Shiu and Bleecker, 2003; Galindo‐Trigo et al., 2016). Analysis of sequences of Populus proteins containing CBM57 revealed that the majority have a kinase domain and that many are homologous to Catharantus roseus RLK1‐like (CrRLK1L) kinases involved in cell wall integrity sensing, polar growth and responses to stresses, such as AtFERRONIA and AtHERKULES1 (Engelsdorf and Hamann, 2014). Protein families with CBM57 were expanded in Populus compared with A. thaliana (Table 1). The evolutionary basis for this expansion is unknown but could be related to more diversified stress reactions in Populus as a result of its perennial lifestyle.

Certain CBMs were observed within Populus GH families (Tables 1 and S7). The most frequently observed combinations were CBM43–GH17 (callose binding domain–glucan endo‐1,3‐β‐glucosidase), CBM18–GH19 (chitin binding domain–chitinase), CBM48–GH13 (glycogen binding domain–amylase) and CBM22–GH10 (xylan binding domain–xylanase). The only GT with a CBM observed in Populus was PtStSy6 from GT5 with three repeats of CBM53, homologous to A. thaliana starch synthase III involved in transitory starch biosynthesis (Valdez et al., 2008).

The CAZy database contains several entries annotated as CBMs in Populus but these lists are incomplete. For example, five out of 15 CBM18‐containing chitinases, one out of eight CBM22‐containing GH10 sequences, eight out of 43 CBM43‐containing endo‐1,3‐β‐glucosidases and nine out of 34 CBM43s containing other proteins were previously identified (http://www.cazy.org; Table 1). Moreover, the smaller CBM families in Populus, such as CBM13, 20, 32, 45 and 53, have not been previously annotated in the CAZy database.

Differential expression of CAZymes in different tissues

Previous CAZyme expression analyses in Populus were based on EST frequencies (Geisler‐Lee et al., 2006) or microarray analysis (Aspeborg et al., 2005). Comparisons of transcriptomes between leaf and developing wood were also made in P. trichocarpa using more sensitive RNA sequencing technology (Hefer et al., 2015). Here, we used RNA sequencing data sets available for aspen (Sundell et al., 2015, 2017; Immanen et al., 2016) to perform comparative expression analyses among larger collections of different tissues, which afforded a more sensitive detection of expressed genes. Indeed, normalized expression values showed that of the 1914 CAZyme and expansin gene models, as many as 1796 (94%) were expressed in at least one of the organs and tissues assayed (Table S9). Ten of these genes (0.5%) were found specifically and highly expressed [organ/tissue specificity score of τ = 1 and variance‐stabilizied (VST) expression ≥1.2], suggesting specialized functions for these genes in these organs/tissues (Table 2).

Table 2.

CAZymes with specific expression pattern in one organ/tissue of aspen and expression value ≥1.2^a

Potri ID	Pt name	CAZy family	Expression (VST)	Specific in:
Potri.002G202100		GH28	1.27	Expanding flowers
Potri.003G223500		EXPN	1.21	Drought‐stressed leaves
Potri.015G040700		AA1	1.29
Potri.008G010700		GH28	1.20	Petiole
Potri.016G107900		AA1	1.25	Roots
Potri.001G351600		CBM43	1.24	Mature seeds
Potri.001G223700		GH1	1.24
Potri.014G082000		EXPN	2.02	Developing phloem
Potri.011G077400	XTH10	GH16	2.10
Potri.005G054000		CBM18‐EXPN	2.00	Developing xylem

^aExpression data from Sundell et al. (2015) and Immanen et al. (2016). All organs/tissues considered as listed in Table S9.

To functionally characterize CAZymes during wood biosynthesis, we examined their expression pattern in the AspWood database (http://aspwood.popgenie.org/aspwood-v3.0/), which represents a high‐spatial‐resolution transcript analysis of developing aspen wood (Sundell et al., 2017). A total of 1187 CAZyme models (62%) were found expressed in wood‐forming tissues (Table S10), the majority of which exhibited defined patterns of expression in developing wood, peaking at different stages of wood formation (Figure 1). These patterns indicate that certain sets of CAZymes have specific functions during wood development, which are separated along the pseudotime developmental program, assayed from the cambium to mature xylem, within these cryosection series. The phloem tissue cluster (as defined in Figure 1) contained the largest number of CAZyme genes, which were linked to cell wall biosynthesis, defense or phloem assimilate transport, as expected. The second largest cluster was associated with the cambium–radial expansion zone, reflecting the diverse metabolic activities required for intense primary cell wall biosynthesis and modification. The primary to secondary wall transition zone was another developmental zone where a large number of CAZymes peaked, most likely in connection with the reorganization of the cell wall biosynthetic machinery.

Figure 1.

Heat map of CAZyme expression patterns showing that the majority of CAZymes expressed in wood‐forming tissues of aspen (1187 genes) have maximum expression (shown as red color) at specific wood developmental stages, defined by the position (section number) on the bottom. Different developmental zones were defined based on the expression of marker genes (Sundell et al., 2017). The expression profiles were sorted by tissue specificity and maximum expression in corresponding tissue. Abbreviations: CA–RE, cambium and radial expansion zone; PW–SW, primary to secondary wall transition; SW, secondary wall formation zone. Data analyzed from AspWood database (http://aspwood.popgenie.org/aspwood-v3.0/).

Subsequent analysis of each cluster composition with regards to different CAZyme families revealed remarkable variation among different wood developmental zones, with different groups of CAZymes dominating in these zones (Figure S2; Table S11). The phloem cluster contained the highest share of CMBs, the cambium–radial expansion zone was characterized by the highest proportion of expansins and PLs, the primary to secondary wall transition zone had the highest proportion of AAs and CEs, the secondary wall formation zone had the highest proportion of GTs and the maturation zone had the highest proportion of GHs (Figure S2; Table S11). Among the GTs, GT1 and GT4 (sucrose synthase) dominated in the phloem, GT2 (largely represented by CSLD genes), GT31, GT57, GT66 and GT75 (UDP‐arabinose mutases) had highest representation in the cambium and radial expansion zone, GT8 (represented mainly by galacturonyl transferases involved in pectin and xylan biosynthesis) dominated during the primary to secondary wall biosynthesis transition and GT43 (involved in xylan biosynthesis) was most prominently represented in the secondary wall formation zone (Figure S2; Table S11). Within the GHs, phloem had the highest share of GH1, GH5 (GH5_11 and GH5_14) and GH14 (β‐amylases), the cambium and radial expansion zone showed the highest abundance of GH16 (XTH), GH9 (cellulases) and GH36, the primary to secondary wall transition zone was most abundant in GH3, GH38 and GH47, the secondary wall formation zone was most abundant in GH28 (polygalacturonases) and the maturation zone ‐ in GH17 (β‐1,3,‐glucanases) (Figure S2; Table S11). This highlights the diversity of CAZyme functions during different stages of xylogenesis.

The cluster analysis (Figure S2; Table S11) provided some insights into the role of the recently annotated CAZyme gene families in Populus. For example, GH152 was abundant in the phloem and cambium–radial expansion zones, consistent with its proposed role in callose metabolism (Trudel et al., 1998; Sakamoto et al., 2006). CBM57 (malectin domain‐receptor‐like kinases) was also most abundantly represented in the same tissues, consistent with the suggested roles in signaling and stress responses (Engelsdorf and Hamann, 2014). Most CE6 members were expressed in the secondary wall formation zone and the AA6 family was most abundant at the primary to secondary transition, which supports the roles of these families in secondary wall biosynthesis. GT106 (Takenaka et al., 2018) had numerous members from different clades reaching high abundance in several xylem cell developmental zones, pointing to their importance at different stages of xylogenesis.

Classification of CAZymes into functional groups related to starch and cell wall biosynthesis

We classified Populus CAZymes into different metabolic groups based on their functional and biochemical characterization or their similarity to enzymes characterized in other plant species (Table S12). Furthermore, as most biochemically and functionally characterized genes were from A. thaliana, we determined their likely orthologs in Populus by phylogenetic analysis, and in some cases proposed their names based on the well‐studied A. thaliana genes (Figure S1; Tables S1–S7). This approach assumes functional conservation for well‐defined clades and although this is generally a valid assumption, there could be exceptional cases of neofunctionalization in one species, especially when linked to disproportionate clade expansion. The clade information as well as highest sequence homology score data are included in the Table S12. The functional categories were broadly classified as starch and sugar related (sucrose, raffinose, stachyose, galactinol and trehalose), cell wall polymer related, including AGPs/extensins, callose, cellulose, xylan, mannan, pectin, xyloglucan and lignin‐related categories. Some enzymes that have a role in more than one type of polymer biosynthesis/modification were separately classified as pectin/xylan, etc. Genes belonging to the same families as the characterized CAZymes but to different clades were classified as ‘uncharacterized’ (Table S12).

Variation of CAZyme transcriptome related to different cell wall polymers during wood formation reflects their biosynthesis pattern

To provide support for our functional annotations, we subsequently analyzed the composition of transcriptomes related to the biosynthesis and degradation of different cell wall components in different zones of developing wood, sampled at high spatial resolution (Sundell et al., 2017), and compared this with changes in cell wall sugar composition and in lignin content during wood development. The CAZymes were clustered according to their maximal expression values as preferentially expressed in different developmental zones, shown in Figure 1, and the number of genes categorized into different metabolic activities were compared among these different clusters. The transcriptome analysis revealed distinct variability across the wood developing zones in both the number of genes with highest expression in a particular zone and in their metabolic functions (Figure 2a; Table S13). Polysaccharide degradation/modification‐related genes were mostly clustered in the phloem and cambium–radial expansion zones. The phloem cluster was relatively rich in starch and sugar degradation‐related genes, whereas in the cambium–radial expansion zone, xyloglucan degradation/modification‐related genes dominated. The primary to secondary wall transition zone was especially abundant in pectin degradation/modification‐related genes, the secondary wall zone was abundant in mannan degradation‐related genes, whereas the maturation zone transcriptome, corresponding mostly to the live xylem parenchyma cells, had the largest share of callose degradation‐related genes. Biosynthesis‐related genes were most abundant in the primary to secondary wall transition zone (Figure 2a). Phloem was enriched in sugar, callose and mannan biosynthetic genes, the cambium–radial expansion zone was enriched in pectin biosynthesis‐related genes, the primary to secondary wall transition zone had the largest share of lignin‐biosynthetic CAZy genes, and the secondary wall zone was enriched in xylan‐related genes. The maturation zone, where mostly parenchyma cells contributed to the transcriptome, was enriched in pectin and starch biosynthesis‐related genes. Pectin constitutes the main part of the protective layer formed by parenchyma cells after secondary wall deposition (Mellerowicz et al., 2001).

Figure 2.

Variability in cell wall composition and in the CAZyme transcriptome related to the biosynthesis and degradation of different polymers across wood developmental zones. (a) Count of gene models associated with specific wood developmental zones (bar graph) and the composition of each group according to predicted functions (pie charts). Data based on Sundell et al. (2017). Gene metabolic classification as listed in Table S12. ‘Sugar’ category includes enzymes related to sucrose, raffinose, stachyose, galactinol and trehalose metabolism. Abbreviations: CA–RE, cambium–radial expansion zone; PW–SW, primary to secondary wall transition; SW, secondary wall formation zone. (b and c) Analysis of cell wall composition in different samples of developing secondary phloem and xylem by alditol acetates (excluding Glc) and uronic acid (UA) contents (b) and Klason lignin content (c). Cell wall samples were obtained by the sequential scraping of bark and wood surfaces exposed after peeling the bark. Alignment between transcriptome and cell wall samplings is shown by the gray shadow. Data in (b) and (c) are mean values from n = 3 technical replicates ± SE.

To reveal whether the observed changes in the transcriptome corresponded to cell wall composition, samples across the developmental gradient of secondary phloem and secondary xylem were obtained by splitting the bark and wood in actively growing aspen trees and sequentially scraping the tissues on both exposed sides. The monosaccharide composition of trifluoroacetic acid (TFA)‐ hydrolyzed polymers and lignin were analyzed in these samples. These samples were aligned with the transcriptome series (shown by gray shading in Figure 2b,c) based on the bark‐splitting site in the radial expansion zone (Gray‐Mitsumune et al., 2004). A shift in monosaccharide composition between primary (radial expansion and outward samples) and secondary (early SW and mature wood) walled samples was clearly observed (Figure 2b), with the primary walled samples having a high content of pectin‐related sugars, arabinose and uronic acids, whereas the secondary walled xylem samples were dominated by xylose, reflecting their high xylan content. The mannose content was relatively constant in all samples. These data agree with published knowledge on the composition of primary and secondary wall layers in hardwoods (Mellerowicz et al., 2001; Mellerowicz and Gorshkova, 2012) and with observed transcriptome variability regarding the pectin and xylan biosynthesis‐related genes (Figure 2a).

The lignin content showed a small peak in the young phloem sample and it increased largely in mature xylem (Figure 3c). The phloem peak identifies the zone of phloem fiber differentiation and is supported by transcript data for lignin‐ and many secondary wall‐related genes (Sundell et al., 2017).

Figure 3.

The co‐expression networks of CAZymes in wood‐forming aspen tissues (http://aspwood.popgenie.org/aspwood-v3.0/). The CAZyme gene co‐expression at a context likelihood of relatedness threshold of 5 or higher was extracted as a network from popgenie (http://popgenie.org/exnet) and visualized in cytoscape 3.4.0. (a) The genes colored by developmental zones where they have peak expression as shown in Fig. 1. (b) The same network colored by the CAZyme metabolic function. ‘Uncharacterized’ class includes members of families with probable function in cell wall biosynthesis. The ‘Sugar’ category includes enzymes related to sucrose, raffinose, stachyose, galactinol and trehalose metabolism. Lists of all genes in the networks are given in Table S14. The guide genes used in the primary wall forming zone are marked by hexagons and those of the secondary wall are marked by diamonds. The first neighbors of the guide genes are listed in Tables S15 and S16.

Uncharacterized CAZymes involved in wood biosynthesis: construction of CAZyme‐based co‐expression networks in wood‐forming tissues

The co‐expression of uncharacterized CAZyme genes with the already known genes in wood‐forming tissues may reveal novel genes involved in wood formation. A total of 471 CAZyme genes formed a co‐expression network, which included many uncharacterized genes with putative functions in cell wall biosynthesis and in other cellular processes during wood formation (Table S14). We have colored the genes in the network by their expression cluster (Figure 3a) and by their metabolic function (Figure 3b). The network clearly illustrates the existence of subnetworks related to different stages of xylem cell differentiation (Figure 3a) and, in a few cases, related to the metabolism of different compounds (Figure 3b). The metabolism of different compounds can be illustrated by the separation of lignin‐ and carbohydrate‐related networks during secondary wall formation, suggesting that these metabolic processes have distinct spatiotemporal regulation.

To identify more candidates for wood cell wall biosynthesis and modification, we have analyzed the closest neighbors of some known cell wall‐related CAZymes using a ‘guide genes’ approach (Aoki et al., 2007) separately for the primary wall‐ and secondary wall‐related networks (Tables S15 and S16, respectively).

Primary wall CAZyme networks

Within the primary wall network, we focused on the first neighbors of the cellulose, pectin and xyloglucan‐related genes that were used as ‘guides’ (Figure 3; Table S15), as the primary wall of aspen xylem contains mostly these components (Mellerowicz et al., 2001). The guide genes included the primary wall CesA PtCesA3‐D (Kumar et al., 2009), the Populus ortholog of AtGAUT7 encoding homogalacturonan α‐1,4‐galacturonosyl transferase (Atmodjo et al., 2011) PtGAUT7-B, the Populus ortholog of AtRGXT1 encoding RG‐II‐α‐1,3‐xylosyl transferase (Egelund et al., 2006; Liu et al., 2011) PtRGXT1-A, and the Populus ortholog of AtXXT3 encoding a candidate xyloglucan α‐1,6‐xylosyl transferase (Vuttipongchaikij et al., 2012) PtXXT3-B/PtGT34B. The obtained cellulose–pectin–xyloglucan network (Table S15) included several putative xyloglucan and pectin biosynthesis‐related genes, as identified by the phylogenetic analysis (Figure S1), such as PtXXT1‐A, PtGAUT9‐A, PtGAUT13‐A, GATL11‐B, two uncharacterized GT47 members from the clade of AtARAD1 involved in pectic arabinan biosynthesis (Potri.T071700 and Potri.019G086800; Harholt et al., 2012), and putative polygalacturonases from GH28 (Potri.016G051200 and Potri.016G054800) (Table S15). Interestingly, the primary wall cellulose–pectin–xyloglucan network included two GT106 members (Table S15), one from the RGI‐I:rhamnosyltransferase (RRT) clade (Potri.015G048100; Takenaka et al., 2018) and another (Potri.003G062600) from a different clade that has not yet been characterized (Figure S1), indicating the involvement of these GT106 members in xylem primary wall biosynthesis. The network also included a GT29 member Potri.014G145400 (Table S15), similar to AtSIA1/AtMGP2 and AtSIA2 transferases that have been proposed to be involved in transferring of DHA or KDO to the RG‐II backbone (Deng et al., 2010; Dumont et al., 2014). Another co‐expressed gene was Potri.019G082200 from GT10. In A. thaliana, Golgi localized GT10 α‐1,3‐or α‐1,4‐fucosyltransferases have been proposed to function in N‐glycan biosynthesis (Both et al., 2011; Rips et al., 2017). Our data suggest a role for the GT10 family in primary cell wall biosynthesis in the xylem. Other genes of this network belong to families GT14, GT31 and GT77, with putative functions in primary wall extensins and AGP biosynthesis (Gille et al., 2013; Knoch et al., 2013; Basu et al., 2015) (Table S15).

The network of the primary wall xylan biosynthetic gene PtGT43E (Ratke et al., 2015, 2018) was distinct from that of the cellulose–pectin–xyloglucan biosynthesis network, and shared some genes with the network of α‐expansin Pt EXPA1 (Gray‐Mitsumune et al., 2008) and pectate lyase Pt PL1‐27 (Biswal et al., 2014), the key expansion markers for xylem cells (Sundell et al., 2017) (Table S15; Figure 3B). This network included additional expansin and PL1 genes, XTH, PL4 members, xylan degradation‐related genes PtXYN10E and PtBXYL2, a cellulase‐encoding gene PtGH9B3 (Takahashi et al., 2009) and a putative mannanase‐encoding gene PtMAN7 (Table S15), which are all likely to be involved in xylem cell expansion. This extends the list of known cell wall modifying enzymes with putative function in xylem cell expansion, and suggests that all classes of polymers are modified during this process. Interestingly, the network also included three members of the AA1 family of the SKU clade that has not yet been characterized in the context of wood development (Figure S1; Table S15). The clade founding the AtSKU5 gene has been implicated in directional growth of roots (Sedbrook et al., 2002), suggesting a role for these Populus SKU5‐like members in wood growth.

Secondary wall CAZyme networks

The secondary wall CAZyme networks included many genes of proven or putative function in the metabolism of xylan, AGP/extensin, cellulose, mannan, lignin and pectin (Figure 3; Table S16). To identify novel candidate genes involved in secondary wall biosynthesis, we analyzed first‐order neighbors of genes that have been shown to be involved in the biosynthesis of: secondary wall cellulose – PtCesA7‐A (Kumar et al., 2009; Song et al., 2010); secondary wall xylan – PtGT43B and PtGT43C, involved in xylan backbone biosynthesis (Lee et al., 2011; Ratke et al., 2018); and PtGT47C, a homolog of AtFRA8, involved in the reducing end sequence biosynthesis in xylan (Lee et al., 2009). The neighbors of PtCesA7‐A, PtGT43B and PtGT47C largely overlapped and included several known genes encoding the components of the secondary wall cellulose synthase complex, such as CesAs (PtCesA4, PtCesA7‐A, PtCesA8‐A and PtCesA8‐B; Song et al., 2010), cellulase PtCel9A1/PtKOR1, which is required for cellulose biosynthesis (Nicol et al., 1998; Szyjanowicz et al., 2004) and cellulose crystallinity regulation (Takahashi et al., 2009; Maloney and Mansfield, 2010), PtGH19A ‐ the homolog of At CTL/ At POM1/ At ELP1, which is essential for cellulose biosynthesis but of unknown molecular function (Sánchez‐Rodríguez et al., 2012), and the sucrose synthase PtSUS2 (Gerber et al., 2014). All these proteins except PtGH19A were found to be co‐immunoprecipitated with secondary wall CesAs in Populus, suggesting their connection to the cellulose synthase complex (Song et al., 2010) and supporting the rationale of using the network analysis for finding additional components of secondary wall biosynthesis. Moreover, we have identified most of the genes with proven or proposed function in secondary wall xylan biosynthesis (Rennie and Scheller, 2014). These included the genes encoding proteins of the xylan synthase complex (Jiang et al., 2016; Zeng et al., 2016): the pair of orthologs to AtIRX10 (Jensen et al., 2014), PtGT47A-1 and PtGT47A-2, one of the pair of orthologs to AtIRX10L (Wu et al., 2009; Mortimer et al., 2015) PtGT47D-1 (Figure S1; Table S12), and AtIRX14 clade member PtGT43D (Ratke et al., 2015, 2018) (Table S16). Thus, similar to Arabidopsis, the xylan:xylosyl transferase activities in secondary walled xylem cells in poplar appear to be redundantly encoded by homologs to AtIRX10 and AtIRX10L (Wu et al., 2009). Furthermore, we identified genes encoding enzymes involved in xylan reducing end biosynthesis in the secondary wall network: PtGAUT12‐A/PtGT8D‐1 and PtGAUT12‐B/PtGT8D‐2 (Li et al., 2011b; Biswal et al., 2015), a pair orthologous to AtIRX8 (Persson et al., 2007); and PtGATL1‐A and PtGATL1‐B, a pair orthologous to AtPARVUS (Kong et al., 2009) (Figure S1; Table S12). The network also included genes encoding putative glucuronosyl transferases, PtGUX1‐A and PtGUX1‐B, a pair orthologous to AtGUX1 (Figure S1; Table S12), and known to be involved in the generation of GlcA decorations on the xylan backbone with an even spacing pattern, forming the major xylan domain of secondary walls (Mortimer et al., 2010; Bromley et al., 2013). Xylan transglycosylase PtXYN10A (Derba‐Maceluch et al., 2015) was also a part of the secondary wall CAZyme network.

Mannan‐related genes found in the secondary wall network included the glucomannan synthase encoding gene PtCSLA1/PtGT2A (Suzuki et al., 2006) and the putative endo‐mannanase gene PtMAN4 (Yuan et al., 2007; Zhao et al., 2013a).

The uncharacterized candidates among the CAZymes identified in the secondary wall network included the CE6 member Potri.014G022600 with putative xylan acetyl esterase function, three members of GT106, one of which was from the clade of RG‐I:rhamnosyl transferases (Takenaka et al., 2018), putative pectin modifying genes from families CE13, PL4 and GH28, and putative AGP synthesizing members of the GT14 and GT31 families (Table S16). The network also included nine AA1 genes belonging to the laccase clade (Figure S1), homologous to AtLAC4/AtIRX12, AtLAC11, AtLAC13 and AtLAC17, of which AtLAC4, ‐11 and ‐17 are known to function in xylem cell lignification (Berthet et al., 2011; Zhao et al., 2013b).

Although several CAZymes known to be involved in secondary wall xylan biosynthesis were co‐expressed as close neighbors of secondary wall CesAs, PtGT43C neighbors did not include secondary wall CesAs except PtCesA8‐A, and did not extensively overlap with the secondary wall xylan network (Figure 3B; Table S16). Instead, this subnetwork included some members of the primary wall CAZyme network, such as two GT106 genes, a putative xyloglucan xylosyl transferase PtXXT3‐B/PtGT34B, a putative homogalacturonan synthase PtGAUT7‐B, and two uncharacterized GT47 members similar to AtARAD1 (Tables S15 and S16). This agrees with a proposal that PtGT43C participates in both the primary and the secondary wall xylan biosynthetic complexes (Ratke et al., 2015, 2018; Sundell et al., 2017). Thus, the PtGT43C subnetwork is expected to include other genes with functions in primary and secondary walls. We identified several genes with putative AGP/extensin biosynthesis functions from families GT14 and GT31 in this subnetwork, as well as genes from uncharacterized CAZyme families: for example, AA6 or GH47 (Table S16).

Associations of CAZymes with other genes involved in wood secondary wall biosynthesis

To identify more co‐expressed genes and their regulators involved in secondary wall biosynthesis in developing xylem, we extended the co‐expression analyses to all genes of P. trichocarpa, using the same stringency (threshold 5) as in CAZyme networks. Cellulose, xylan and glucomannan are the main polysaccharides of secondary wall layers in aspen, and hence we used CAZyme genes involved in the biosynthesis of these polymers as guide genes, including the prominent members of the identified CAZyme network associated with secondary wall biosynthesis (Table S16). By using this approach, we identified 354 gene models co‐expressed with one or more of secondary wall guide genes (Table S17).

As expected, the network included the previously identified CAZymes with known or putative functions in secondary wall cellulose, xylan and glucomannan biosynthesis. In aspen wood, all matrix polysaccharides are acetylated, which affects their water solubility, ability to undergo enzymatic modification of degradation, and interactions with cellulose and other polymers (Pawar et al., 2013; Busse‐Wicher et al., 2014). Accordingly, the acetylation‐related genes were a prominent group in the secondary wall network. They included PtRWA‐A (Pawar et al., 2017), several members of the TBL family of acetyltransferases homologous to AtTBL3, ‐29, ‐33 and ‐34 (Xiong et al., 2013; Yuan et al., 2013, 2016a,2016b,2016c; Urbanowicz et al., 2014), as well as AtAXY9 (Schultink et al., 2015) homolog Potri.017G139800 (Table S17, cf. acetylation category).

The glucuronic acid side chain of xylan in woody species, including aspen, is methylated at O‐4 (Teleman et al., 2000), and this methylation is essential for wood growth and secondary wall formation (Song et al., 2014). It is generated in the Golgi with the help of glucuronoxylan methyl transferases (GXMTs) found in clade A of the DUF579 family (Lee et al., 2012; Urbanowicz et al., 2012). So far, GXMT activity has been identified in four members of the Populus DUF579 gene family, PtGXMT1/PtDUF579‐2, PtGXMT2/PtDUF579‐1, PtGXMT3/PtDUF579‐3 and PtGXMT4/PtDUF579‐4 (Song et al., 2014; Yuan et al., 2014), with the first three being part of the secondary wall network (Table S17, cf. DUF579 category). Clade B of the DUF579 family contains a pair of AtIRX15 and AtIRX15‐L genes that are required for glucuronoxylan biosynthesis in secondary cell walls (Brown et al., 2011; Jensen et al., 2011), their orthologous pair PtDUF579‐10 and PtDUF579‐9 (Song et al., 2014) were also a part of the secondary wall network (Table S17, cf. DUF579 category). Interestingly, PtDUF579‐9 is expressed with PtGH43B, whereas PtDUF579‐10 is expressed as a first neighbor of PtGT43C (Table S17). These two expression networks formed separate clusters with very little overlap (Figure 4a), and we propose that the former is associated strictly with the secondary wall formation and the latter includes genes acting at both primary and secondary wall formation stages (Ratke et al., 2015, 2018).

Figure 4.

The difference in neighborhoods of genes encoding proteins of secondary wall xylan synthase complex Pt GT43B and Pt GT43C (a), and secondary wall cellulose synthase complex PtCesA8‐A and PtCesA8‐B (b). The co‐expressed genes were selected at threshold = 5 (http://aspwood.popgenie.org/aspwood-v3.0/). The blue nodes are unique neighbors of Pt GT43C (in a) and PtCesA8‐B (in b) and the orange nodes are unique for Pt GT43B (in a) and PtCesA8‐A (in b). The pink nodes are neighbors in common for both genes in (a) and (b). The transcription factors are marked by rectangles. Unless otherwise indicated, the Populus gene names are used in the figures.

The fasciclin‐like arabinogalactan proteins (FLAs) play an important role in cell wall architecture in A. thaliana and Populus, mediating wall mechanical properties (Johnson et al., 2003; Andersson‐Gunnerås et al., 2006; Wang et al., 2015). We observed several homologs of AtFLA17 and AtFLA11 in the secondary wall network (Table S17, cf. AGP category). Their presence, along with the presence of a lysine‐rich arabinogalactan protein similar to AtAGP19, is matched with the abundance of CAZymes with proposed function in AGP glycosylation (Table S16).

The secondary wall network had a prominent representation of proteins involved in vesicle formation and movement, and in rosette movement (Table S17, cf. vesicle, movement category). The identified genes are good candidates for the biotechnological regulation of secondary wall formation, as the regulation of vesicle and cellulose synthase complex movement is likely to limit the biosynthesis of the secondary wall (McFarlane et al., 2014; Wang et al., 2016).

Another very prominent group of the secondary wall network were the signaling‐ and protein modification‐related genes, including genes encoding receptor‐like kinases with a lectin domain (Table S17, cf. ‘signaling’ category). Several of these proteins are known as receptors involved in cell wall integrity (CWI) signaling that perceive cell wall damage and activate diverse responses, including gene expression (Engelsdorf and Hamann, 2014). CWI was described in primary walled cells, but whether equivalent CWI signaling exists in cells depositing secondary walls is currently under debate because no expected changes in gene expression were detected in A. thaliana mutants affected in secondary wall xylan biosynthesis (Faria‐Blanc et al., 2018). The mutants in secondary wall CesA (Hernandez‐Blanco et al., 2007) and secondary wall acetylation (Pawar et al., 2016) exhibited altered expression of resistance genes, however, as well as increased resistance to pathogens, which were assumed to be triggered by CWI. Moreover, Ratke et al. (2018) reported altered gene expression and increased growth in aspen with reduced secondary wall xylan biosynthetic GT43 genes, and suggested that these changes were mediated by CWI in secondary walled cells. Our finding of several receptor‐like kinases in the secondary wall network (Table S17) provides strong candidates for secondary CWI sensing. Interestingly, the dominant repressors of the double mutant fei1 fei2 (impaired in CWI) have been recently identified as SHOU4 and SHOU4L in A. thaliana (Polko et al., 2018), and the secondary wall network contained three homologs of these genes associated with PtGT43C (Figure 4a; Table S17, cf. cell wall category), strengthening the hypothesis of the secondary CWI pathway.

More attention has been given to the transcription factors regulating the onset of secondary wall development (Ohtani et al., 2011; Zhong et al., 2011; Taylor‐Teeples et al., 2015), as they can offer direct genetic approaches to improve the productivity of trees. We have identified several well‐known transcription factors as co‐expressed neighbors with different secondary wall biosynthetic CAZyme genes (Table 3). As we observed clear differences in networks of PtGT43C and PtGT43B, confirming the previous conclusions that PtGT43C functions in both primary and secondary xylan biosynthesis, whereas PtGT43B is specifically involved in secondary wall xylan biosynthesis (Ratke et al., 2015, 2018), we analyzed the transcription factors in these networks in order to reveal the nature of their regulons (Figure 4a). The secondary wall‐associated NAC domain (SND) and vascular‐related NAC domain (VND) proteins are known as master switchers of secondary wall formation in Populus (Ohtani et al., 2011; Zhong et al., 2011; Li et al., 2012). The overexpression of PtSND1‐B2/PtWND2B resulted in the ectopic deposition of secondary walls along with enhanced expression of the genes involved in the biosynthesis of wood components, cellulose, xylan and lignin (Zhong et al., 2011; Li et al., 2012). We identified PtSND1‐B2/PtWND2B co‐expressed as a close neighbor of PtGT43B (Figure 4a; Table 3), in agreement with the previous reports of PtSND1‐B2/PtWND2B activating the PtGT43B promoter (Ratke et al., 2015). Some of the important primary targets of PtSND1‐ B2/PtWND2B, such as PtKNAT7, PtNAC156, PtBLH3, PtMYB128 and PtMYB10 (Zhong et al., 2011), have also been found associated with secondary wall CAZymes (Table 3). Of these, PtNAC156, PtBLH3 and PtMYB128 were associated with PtGT43B, but not with PtGT43C (Figure 4a). PtGT43B was also uniquely co‐expressed with an ethylene response factor (ERF) transcription factor PtERF119 homologous to AtSHINE3 (AtSHN3), which was recently found as an important hub transcription factor in secondary wall forming aspen xylem (Seyfferth et al., 2018). Overexpression of the closely related Populus SHN2/ERF118 gene induced cellulose and hemicellulose and suppressed lignin biosynthesis in Nicotiana tabacum (tobacco; Liu et al., 2017). The PtGT43C network included genes encoding transcription factors that have not been studied in the context of secondary growth, such as homologs of the GATA family gene AtGATA9 and an ARF family gene AtARF7 implicated in auxin‐induced lateral root growth and tropism responses (Goh et al., 2012). These and other transcription factors of unknown function found in our analysis (Table 3) could represent targets for tree improvement.

Table 3.

Transcription factors co‐regulated with genes encoding enzymes synthesizing cellulose, mannan and xylan in secondary walls in the developing wood of aspen (AspWood, http://aspwood.popgenie.org/aspwood-v3.0/)

Potri. ID	Potri name	AT ID	AT name	TF class	CesA7‐A Potri.006G181900	CesA8‐A Potri.011G069600	CesA8‐B Potri.004G059600	GT43B Potri.016G086400	GT43C Potri.007G047500	GT47A‐1 Potri.001G068100	GATL1‐B Potri.002G132900	GATL1‐A Potri.014G040300	GT47C Potri.009G006500	CSLA1 Potri.008G026400	GUX1‐A Potri.007G107200	GUX1‐B Potri.005G061600
001G099800	MYB10	1G63910	MYB103	MYB	+		+						+
001G112200	KNAT7	1G62990	IXR11, KNAT7	TALE	+		+						+
001G258400				C3H	+		+			+	+					+
002G119400		3G49940	LBD38	LBD	+		+			+	+					+
002G257800		4G28610	PHR1	G2‐like	+
006G253800		5G25390	SHN3	ERF	+			+		+	+					+
007G135300	NAC156	4G28500	ANAC073, NAC073, SND2	NAC	+	+	+	+		+	+	+				+
008G117500		1G68670	HHO2	G2‐like	+	+	+									+
009G045900		5G60440	AGL62	M‐type_MADS	+		+			+	+	+				+
017G016700	NAC154	4G28500	ANAC073, NAC073, SND2	NAC	+		+				+		+	+		+
002G031000	BLH3	2G16400	BLH7	TALE		+		+
002G178700	WND2B, SND1‐B2	2G46770	ANAC043, EMB2301, NST1	NAC				+
003G132000	MYB128	1G63910	MYB103	MYB				+
006G138500		5G20730	ARF7,BIP,IAA21,‐23,‐25, MSG1,NPH4,TIR5	ARF					–
008G038900		4G32890	GATA9	GATA					+
001G141100		5G46690	bHLH071	bHLH							+
012G055700		1G73830	BEE3	bHLH												+
005G221400		2G42410	ZFP11	C2H2											+

+, positive; –, negative correlation

We also noted subtle differences between the CAZyme networks of PtCesA8A and those of other secondary wall CesAs. In A. thaliana, AtCesA4, ‐7 and ‐8 are found to be co‐expressed, as their encoded proteins form the secondary wall cellulose synthase complex in equimolar ratios (Hill et al., 2014). The situation is more complex in Populus, because there are two paralogs of each AtCesA7 and AtCesA8 (Kumar et al., 2009), creating the possibility of neofunctionalization for one of these duplicated members. We suspected that this could be the case for the PtCes8‐A/B pair as their expression in aspen wood‐forming tissues diverged during the later stages of xylogenesis (Sundell et al., 2017). The co‐expressed neighbors between PtCesA8‐A and PtCesA8‐B had significant similarity for cellulose biosynthetic proteins (for example, PtCesA4, PtCes7‐A, PtCes7‐B and PtGH19A), but showed differences for transcription factors and signaling‐related proteins (Figure 4b; Tables 3 and S17). They both shared PtNAC156 (Zhong et al., 2011) and Potri.008G117500, a MYB‐like gene with unknown function homologous to AtHHO2, controlling phosphate homeostasis (Nagarajan et al., 2016), but PtCesA8‐A was uniquely associated with PtBLH3, a transcription factor downstream of PtSND1‐B2/PtWND2B (Zhong et al., 2011), and a SAUR‐like gene, suggesting the involvement of IAA in the regulation of this network. The PtCesA8‐A network also included secondary xylan biosynthetic genes strictly expressed during secondary wall formation, PtGT43A and PtGT43B (Ratke et al., 2015), suggesting a secondary wall function for PtCesA8‐A (Figure 4b). The unique neighbors of PtCesA8‐B included other secondary wall‐controlling transcription factors, including PtKNAT7, PtMYB10 and PtNAC154 (Zhong et al., 2011), as well as a MADS‐box gene homologous to AGAMOUS LIKE 62 (AtAGL62), which is known to regulate the cellularization of the endosperm in A. thaliana (Kang et al., 2008) (Figure 4b; Table 3). The presence of the AtETO1 homolog in this subnetwork suggests its negative correlation with ethylene signaling; AtETO1 is known to target ACC synthases that are rate‐limiting enzymes in ethylene biosynthesis for proteolysis (Christians et al., 2009). Taken together, these observed differences in the neighborhoods of PtCes8‐A and PtCesA8‐B suggest some additional function(s) for PtCesA8‐B in wood development beyond a core secondary wall CesA function. Indeed, PtCesA8‐B was found to be the dominant CesA isoform expressed in tension wood of aspen, possibly even forming homomeric cellulose synthase complexes in this tissue (Zhang et al., 2018).

Conclusion

This study provides a census of all CAZymes and expansin‐related proteins in P. trichocarpa – a model hardwood species – based on the re‐annotation of gene models of the v3.0 genome assembly. The abundance and diversity of CAZymes in P. trichocarpa is compared with those in A. thaliana revealing no differences in the presence of CAZyme families, but considerable differences in the relative size of certain families, suggesting their adaptive regulation. We report 101 CAZyme families, 18 of which have not been previously annotated in P. trichocarpa, and the updated information on family members with 1914 genes in total annotated as CAZymes, which is a major update from the first annotation based on the v1.0 assembly (Geissler‐Lee et al., 2009). The availability of the AspWood database (http://aspwood.popgenie.org/aspwood-v3.0/) and the aspen expression atlas (Sundell et al., 2015), as well as published aspen RNA‐Seq data (Immanen et al., 2016), provided evidence for the expression for 94% of the gene models, and the expression of 62% of the models in developing wood tissues, and enabled comparative analyses of expression. We used the CAZymes expression profiles during wood development and their relatedness to create corresponding co‐expression networks and to identify regulons composed of CAZymes and other genes. Transcriptional changes occurring during wood development provided several candidates for the biosynthesis of primary and secondary cell wall in P. trichocarpa. Our findings support the existence of two separate regulons for genes involved in secondary wall xylan biosynthesis: one strictly limited to secondary wall formation and another with a broader expression pattern encompassing the cambium and radial expansion zone. Moreover, the data provide support for the diversification of functions of the Populus secondary cellulose synthase paralogs PtCesA8‐A and PtCesA8‐B. The collection of tissue‐specific uncharacterized genes and transcription factors offers a rich source of targets for the future genetic improvement of woody plants.

Experimental procedures

Identification and annotation of CAZymes families in P. trichocarpa v3.0

The protein sequences from 41 335 loci of the P. trichocarpa genome (v3.0) were downloaded from the US Department of Energy (DOE) Joint Genome Institute (https://phytozome.jgi.doe.gov/pz/portal.html#!bulk?org=Org_Ptrichocarpa) and matched to over 1 million curated entries in the CAZy database as of November 2018 (http://www.cazy.org).

Establishment of CAZyme expression matrix for comparative analyses

The high‐resolution RNA‐Seq data sets for the wild and glasshouse grown aspen (Populus tremula L.) are available from the PlantGenIE website (Sundell et al., 2015), and those for glasshouse‐grown hybrid aspen phloem, cambium and developing xylem are detailed by Immanen et al. (2016). The retrieved data were pre‐processed according to our guidelines as described by Delhomme et al. (http://www.epigenesys.eu/en/protocols/bio-informatics/1283-guidelines-for-rna-seq-data-analysis). Briefly, the quality of the raw sequence data was assessed using fastqc (http://www.bioinformatics.babraham.ac.uk/projects/fastqc/). Residual ribosomal RNA (rRNA) contamination was assessed and filtered using sortmerna 2.1 (with settings –log –fastx –sam –num_alignments 1 for both data sets, with the addition of –paired_in for the AspWood data set; Kopylova et al., 2012) using the rRNA sequences provided with sortmerna (rfam‐5s‐database‐id98.fasta, rfam‐5.8s‐database‐id98.fasta, silva‐arc‐16s‐database‐id95.fasta, silva‐bac‐16s‐database‐id85.fasta, silva‐euk‐18s‐database‐id95.fasta, silva‐arc‐23s‐database‐id98.fasta, silva‐bac‐23s‐database‐id98.fasta and silva‐euk‐28s‐database‐id98.fasta). Data were then filtered to remove adapters and trimmed for quality using trimmomatic 0.32 (with settings TruSeq3‐PE‐2.fa:2:30:10 LEADING:3 SLIDINGWINDOW:5:20 MINLEN:50; Bolger et al., 2014). After both filtering steps, fastqc was run again to ensure that no technical artefacts were introduced. Filtered reads were aligned to v3.0 of the P. trichocarpa genome (https://phytozome.jgi.doe.gov/pz/portal.html, November 2018), using star 2.5.2b (with non‐default settings –outSAMstrandField intronMotif –readFilesCommand zcat –outSAMmapqUnique 254 –quantMode TranscriptomeSAM –outFilterMultimapNmax100 –outReadsUnmapped Fastx –chimSegmentMin1 –outSAMtype BAM SortedByCoordinate –outWigType bedGraph –alignIntronMax11000; Dobin et al., 2013). The annotations obtained from the P. trichocarpa v3.0 GFF file were flattened to generate ‘synthetic’ gene models. This synthetic transcript GFF file and the star read alignments were used as input to the HTSeq (Anders et al., 2014) htseq‐count python utility to calculate exon‐based read count values. The htseq‐count utility takes only unique mapping reads into account. Because of their different characteristics (differences in sampling, sequencing instruments, etc.), the data sets were normalized independently. First, the data sets were normalized using variance stabilizing transformation (VST) in r 3.4.0 (R Core Team, 2015) using the bioconductor 3.4 (Gentleman et al., 2004) deseq2 package (v1.16.1; Love et al., 2014). Furthermore, the data from Immanen et al. (2016) were then filtered and samples corresponding to the zones of interest were identified (phloem, cambium and developing xylem) and selected from the three control trees: 4A, 4B and 6B. A likely artefact introduced by decreasing library sizes as samples moved inwards (towards the xylem) was identified and was corrected using a linear modeling approach. Finally, the data from both data sets were merged together using a sample‐based median‐centering approach. The resulting data were subjected to hierarchical clustering and the τ tissue/sample specificity score (Yanai et al., 2004) was calculated, both in r. The whole data analysis was performed using custom r scripts available from our GitHub repository (https://github.com/UPSCb/UPSCb/tree/master/manuscripts/Kumar2018) and the corresponding data are available from our FTP repository (ftp://ftp.plantgenie.org/Publications/Kumar2018).

To identify the poplar orthologs of characterized CAZyme genes of A. thaliana, the protein sequences were aligned using muscle (http://phylogeny.lirmm.fr/phylo_cgi/index.cgi) and phylogenic trees were constructed using the neighbor‐joining (NJ) method of mega 7 (Kumar et al., 2016) in default mode with a bootstrap test of 1000 replicates. Arabidopsis thaliana orthologs of the identified poplar genes were identified by blast (https://www.arabidopsis.org/Blast/).

CAZymes involved in wood biosynthesis

The AspWood high‐spatial‐resolution RNA‐Seq data set was used for the functional analysis of CAZymes in wood formation (Sundell et al., 2017). The identity of wood developmental zones was based on the expression of the marker genes (Sundell et al., 2017). A heat map of CAZyme expression was constructed using the r 3.4.0 (R Core Team 2015) gplots package and the clustering was performed using the Ward.D2 method on Pearson distances. The custom scripts are available from the same GitHub repository described above. CAZyme gene clustering was kept from the published hierarchical clusters (Sundell et al., 2017). To evaluate the CAZymes transcriptome investment in the development of various tissues in wood, we calculated the average VST expression values for all samples covering specified developmental zones in tree 1.

CAZymes networks

Co‐expression networks for all expressed CAZymes were obtained from the AspWood database (http://aspwood.popgenie.org/aspwood-v3.0/) and the corresponding Graphml file was obtained using the exnet tool (http://popgenie.org/exnet) at a Z‐score threshold of 5.0, and is available at ftp://ftp.plantgenie.org/Publications/Kumar2018. The co‐expression networks represented at AspWood were calculated using mutual information (MI) and context likelihood of relatedness (CLR), as explained in Sundell et al. (2017). The resulting Graphml file was further explored in cytoscape 3.4.0 (Shannon et al., 2003) for visualization, coloring and determining the first neighbors of functionally known ‘Guide Genes’ involved in the development of primary and secondary walls of the plant cell (Aoki et al., 2007).

Association of secondary wall‐related CAZymes with other genes

Selected ‘Guide Genes’ were used to extract their corresponding networks containing first neighbors at threshold of 5.0 from AspWood. The similarity and differences in the neighborhoods of CAZymes were visualized using cytoscape 3.4.0.

Data statement

This manuscript is based on publicly available data sets deposited at Phytozome (https://phytozome.jgi.doe.gov/pz/portal.html), CAZy (http://www.cazy.org) and AspWood (http://aspwood.popgenie.org/aspwood-v3.0/; Sundell et al., 2017). The CAZy annotation is provided in supplementary Tables S1–S8.

Cell wall chemical analysis in different developmental zones of secondary xylem and phloem

Stem segments from the basal part of two glasshouse‐grown 6‐month‐old hybrid aspen trees (P. tremula L. × tremuloides Michx.), clone T89, were split into bark and wood core, and sequentially scraped on both exposed surfaces to obtain two sequential developmental stages of wood and secondary phloem differentiation. The rest of the wood core was analyzed as ‘mature wood’. The tissues were freeze‐dried, ground and analyzed for monosaccharide composition by alditol acetates (Englyst and Cummings, 1984) after hydrolysis in 2M TFA, for uronic acid content by a biphenyl assay (Filisetti‐Cozzi and Carpita, 1991), and for Klason lignin content as described previously (Gandla et al., 2015). The developmental trend was similar for the two trees analyzed, and results from one tree are shown in Figure 2 whereas the results for the other tree are shown in Figure S3.

Funding information

We acknowledge VR and Formas grants to E.J.M., the SSF project ValueTree RBP14‐0011, the support from Vinnova (the Swedish Governmental Agency for Innovation Systems) and KAW (The Knut and Alice Wallenberg Foundation), and the Kempe post‐doc stipendium to V.K.

Author contributions

VK performed most data analyses, prepared all figures and tables, and wrote the manuscript with EJM, MH and BH performed CAZyme re‐identification. ND, CM and NRS assisted with data retrieval and transcriptome analyses. P.I. was responsible for cell wall analyses. EJM conceived and coordinated the project, finalized the manuscript with contributions from all co‐authors, and agrees to serve as the author responsible for correspondence.

Conflict of interest

The authors declare no conflicts of interest.

Supporting information

Figure S1. Phylogenetic trees of selected CAZyme families in poplar and A. thaliana.

Figure S2. Variation of CAZymes expression across the wood‐forming zones of aspen.

Figure S3. Variability in cell wall composition across wood developmental zones for the second tree analyzed.

Table S1. List of annotated glycoside hydrolases (GHs) in P. trichocarpa genome v3.0.

Table S2. List of annotated glycosyl transferases (GTs) in P. trichocarpa genome v3.0.

Table S3. List of annotated polysaccharide lyases (PLs) in P. trichocarpa genome v3.0.

Table S4. List of annotated carbohydrate esterases (CEs) in P. trichocarpa genome v3.0.

Table S5. List of annotated auxiliary activities (AAs) in P. trichocarpa genome v3.0.

Table S6. List of annotated expansins (EXPN) in P. trichocarpa genome v3.0.

Table S7. List of annotated carbohydrate binding motifs (CBMs) in P. trichocarpa genome v3.0.

Table S8. Arabidopsis thaliana auxiliary activities (AAs).

Table S9. CAZyme expression matrix for comparative analyses.

Table S10. CAZymes expressed in developing secondary xylem and phloem of aspen.

Table S11. Number of CAZymes per family and per each expression cluster corresponding to wood developmental zones.

Table S12. List of CAZymes with documented and likely functions in cell wall biosynthesis and modification, and in starch and sugar metabolism.

Table S13. Number of CAZymes per each metabolic activities and per each expression cluster corresponding to wood developmental zones.

Table S14. List of genes from CAZymes‐based networks in wood‐forming aspen tissues.

Table S15. List of first neighbors of primary wall‐associated guide genes.

Table S16. List of first neighbors of secondary wall‐associated guide genes.

Table S17. List of first neighbors of secondary wall‐expressed selected CAZymes.