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

Vikash Kumar, Matthieu Hainaut, Nicolas Delhomme, Chanaka

Mannapperuma, Peter Immerzeel, Nathaniel R. Street, Bernard Henrissat,
Ewa J. Mellerowicz

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Vikash Kumar, Matthieu Hainaut, Nicolas Delhomme, Chanaka Mannapperuma, Peter Immerzeel, et al.. Poplar carbohydrate-active enzymes: whole-genome annotation and functional analyses based on RNA expression data. Plant Journal, 2019, 10.1111/tpj.14417. hal-02628965

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Distributed under a Creative Commons Attribution 4.0 International License
The Plant Journal (2019)

doi: 10.1111/tpj.14417

RESOURCE

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

Vikash Kumar<fnref n="1" />, Matthieu Hainaut<fnref n="2" />,<fnref n="3" />, Nicolas Delhomme<fnref n="1" />, Chanaka Mannapperuma<fnref n="4" />, Peter Immerzeel<fnref n="1" />,<fnref n="5" />,

Nathaniel R. Street<fnref n="4" />, Bernard Henrissat<fnref n="2" />,<fnref n="3" /> and Ewa J. Mellerowicz<fnref n="1" />,*

Received 6 December 2018; revised 6 May 2019; accepted 13 May 2019.

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.

Keywords: carbohydrate metabolism, cell wall, comparative genomics, genome sequencing, vegetative development, wood formation.

INTRODUCTION

Carbohydrate-active enzymes (CAZymes) are essential pro-
teins for all life forms (Lombard et al., 2014). They are fun-
damentally important for the synthesis, degradation and
modification of all the glycoconjugates found in nature,
including carbohydrates, glycoproteins, glycolipids and

glycosylated secondary metabolites (Wilson, 2002; Can-
tarel et al., 2009). Plant tissues are particularly rich in com-
plex 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,

<footnote n="1">¹Umeå Plant Science Center, Department of Forest Genetics and Plant Physiology, Swedish University of Agricultural Sciences, Umeå, Sweden,</footnote>

<footnote n="2">²Architecture et Fonction des Macromolécules Biologiques, Centre National de la Recherche Scientifique (CNRS), Aix-Marseille University, Marseille, France,</footnote>

<footnote n="3">³INRA, USC 1408 AFMB, Marseille, France,</footnote>

<footnote n="4">⁴Umeå Plant Science Center, Plant Physiology Department, Umeå University, Umeå, Sweden, and</footnote>

<footnote n="5">⁵Chemical Engineering, Karlstad University, Karlstad 65188, Sweden</footnote>

<footnote>*For correspondence (e-mail ewa.mellerowicz@slu.se).</footnote>
defense, dormancy, signaling and photosynthesis (Cou-
tinho et al., 2003). As a result of high abundance and sus-
tainable 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 simul-
taneously, 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 pro-
tein sequence (Lombard et al., 2014). By adopting this
approach, the CAZymes were previously classified into
four major classes, namely glycosyltransferases (GTs), gly-
coside hydrolases (GHs), polysaccharide lyases (PLs) and
carbohydrate esterases (CEs) (Cantarel et al., 2009).
Recently, the lignin degrading enzymes and the polysac-
charide lytic monooxygenases have been added and classi-
fied as auxiliary activity (AA) families, which encompass a
broad range of enzymes related to lignocellulose conver-
sion (Levasseur et al., 2013). In plants, this group also con-
tains 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, lig-
nin 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 (Gratta-
paglia et al., 2009; Hinchee et al., 2009). As CAZymes are
involved in the formation and modification of this carbohy-
drate matrix (Mellerowicz and Sundberg, 2008), a detailed
study of CAZymes in a woody model species and their spa-
tial interactions during wood cell wall formation is essen-
tial 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 Popu-
lus. 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 anno-
tation (Geisler-Lee et al., 2006) and with the current Ara-
bidopsis thaliana (http://www.cazy.org) annotation.
Furthermore, we have annotated CAZyme families in P. tri-
chocarpa 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.

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

<footnote><sup>a</sup>Families 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.</footnote>

<footnote>* 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).</footnote>
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-β-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 mem-
bers. 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 com-
prises 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 mem-
bers 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 pec-
tins and glucosaminoglycans using a β-elimination mecha-
nism 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 Ca²¹ binding site or other motifs present in PL1_1 (Bai et al., 2017).

Auxiliary activities. The category broadly termed as 'aux-
iliary activities' (AAs) has been introduced to the CAZy
database since the previous Populus genome-wide annota-
tion (Geisler-Lee et al., 2006). It groups together the fami-
lies of redox enzymes and lytic polysaccharide
monooxygenases (LPMOs) involved in polysaccharide,
oligosaccharide and lignin breakdown, lignin polymeriza-
tion 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 pro-
teins 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 consid-
ered 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 reg-
ulation 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 tomen-
tosa (http://www.cazy.org). Closer inspection of P. tri-
chocarpa 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*, con-
tains 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 expan-
sion is unknown but could be related to more diversified
stress reactions in *Populus* as a result of its perennial life-
style.

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 expansion 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-stabilized (VST) expression ≥1.2], suggesting specialized functions for these genes in these organs/tissues (Table 2).

To functionally characterize CAZymes during wood
biosynthesis, we examined their expression pattern in the
AspWood database (http://aspwood.popgenie.org/asp
wood-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
<table><caption>**Table 2** CAZymes with specific expression pattern in one organ/tissue of aspen and expression value ≥1.2ª</caption><thead><tr><th>Potri ID</th><th>Pt name</th><th>CAZy family</th><th>Expression (VST)</th><th>Specific in:</th></tr></thead><tbody><tr><td>Potri.002G202100</td><td></td><td>GH28</td><td>1.27</td><td>Expanding flowers</td></tr><tr><td>Potri.003G223500</td><td></td><td>EXPN</td><td>1.21</td><td>Drought-stressed leaves</td></tr><tr><td>Potri.015G040700</td><td></td><td>AA1</td><td>1.29</td><td></td></tr><tr><td>Potri.008G010700</td><td></td><td>GH28</td><td>1.20</td><td>Petiole</td></tr><tr><td>Potri.016G107900</td><td></td><td>AA1</td><td>1.25</td><td>Roots</td></tr><tr><td>Potri.001G351600</td><td></td><td>CBM43</td><td>1.24</td><td>Mature seeds</td></tr><tr><td>Potri.001G223700</td><td></td><td>GH1</td><td>1.24</td><td></td></tr><tr><td>Potri.014G082000</td><td></td><td>EXPN</td><td>2.02</td><td>Developing phloem</td></tr><tr><td>Potri.011G077400</td><td>XTH10</td><td>GH16</td><td>2.10</td><td></td></tr><tr><td>Potri.005G054000</td><td></td><td>CBM18-EXPN</td><td>2.00</td><td>Developing xylem</td></tr></tbody></table>

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.

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 expansions 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

<figure><img src="image_3.png" /><figcaption>**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 Asp-Wood database (http://aspwood.popgenie.org/aspwood-v3.0/).</figcaption></figure>

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

<footnote>ªExpression data from Sundell *et al.* (2015) and Immanen *et al.* (2016). All organs/tissues considered as listed in Table S9.</footnote>
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 abun-
dant 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) (Fig-
ure 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 expan-
sion 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, consis-
tent 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 pri-
mary to secondary transition, which supports the roles of
these families in secondary wall biosynthesis. GT106 (Tak-
enaka *et al.*, 2018) had numerous members from different
clades reaching high abundance in several xylem cell
developmental zones, pointing to their importance at dif-
ferent 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 charac-
terization or their similarity to enzymes characterized in
other plant species (Table S12). Furthermore, as most bio-
chemically and functionally characterized genes were from
*A. thaliana*, we determined their likely orthologs in *Popu-
lus* 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 func-
tional conservation for well-defined clades and although
this is generally a valid assumption, there could be excep-
tional cases of neofunctionalization in one species, espe-
cially when linked to disproportionate clade expansion.
The clade information as well as highest sequence homol-
ogy score data are included in the Table S12. The func-
tional categories were broadly classified as starch and
sugar related (sucrose, raffinose, stachyose, galactinol and
trehalose), cell wall polymer related, including AGPs/ex-
tensins, callose, cellulose, xylan, mannan, pectin, xyloglu-
can and lignin-related categories. Some enzymes that have
a role in more than one type of polymer biosynthesis/mod-
ification were separately classified as pectin/xylan, etc.
Genes belonging to the same families as the characterized
CAZymes but to different clades were classified as ‘unchar-
acterized’ (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 sub-
sequently 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 compo-
sition 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 tran-
scriptome 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 meta-
bolic functions (Figure 2a; Table S13). Polysaccharide
degradation/modification-related genes were mostly clus-
tered 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-re-
lated genes dominated. The primary to secondary wall
transition zone was especially abundant in pectin degrada-
tion/modification-related genes, the secondary wall zone
was abundant in mannan degradation-related genes,
whereas the maturation zone transcriptome, correspond-
ing mostly to the live xylem parenchyma cells, had the lar-
gest share of callose degradation-related genes.
Biosynthesis-related genes were most abundant in the pri-
mary 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 sec-
ondary 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 tran-
scriptome, was enriched in pectin and starch biosynthesis-
related genes. Pectin constitutes the main part of the pro-
tective layer formed by parenchyma cells after secondary
wall deposition (Mellerowicz et al., 2001).

To reveal whether the observed changes in the transcrip-
tome 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 monosac-
charide composition of trifluoroacetic acid (TFA)- hydro-
lyzed polymers and lignin were analyzed in these samples.
These samples were aligned with the transcriptome series
<figure><img src="image_4.png" /><figcaption>**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.</figcaption></figure>

(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).

## 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
<figure><img src="image_5.png" /><figcaption>**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.</figcaption></figure>

separation of lignin- and carbohydrate-related networks
during secondary wall formation, suggesting that these
metabolic processes have distinct spatiotemporal regula-
tion.

To identify more candidates for wood cell wall biosyn-
thesis 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 biosynthe-
sis. 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 back-
bone (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-fucosyl-
transferases 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 *PtEXPA1* (Gray-Mitsumune et al., 2008) and pectate lyase *PtPL1-27* (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 *AtCTL/AtPOM1/AtELP1*, 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 sec-
ondary 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 subnet-
work included some members of the primary wall CAZyme
network, such as two GT106 genes, a putative xyloglucan
xylosyl transferase PtXXT3-B/PtGT34B, a putative homo-
galacturonan synthase PtGAUT7-B, and two uncharacter-
ized 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 biosyn-
thetic 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 sec-
ondary 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 regula-
tors involved in secondary wall biosynthesis in develop-
ing 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 sec-
ondary wall layers in aspen, and hence we used
CAZyme genes involved in the biosynthesis of these
polymers as guide genes, including the prominent mem-
bers 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 identi-
fied CAZymes with known or putative functions in sec-
ondary 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 acetyl-
transferases 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 spe-
cies, 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 gen-
erated in the Golgi with the help of glucuronoxylan methyl
transferases (GXMTs) found in clade A of the DUF579 fam-
ily (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 pri-
mary and secondary wall formation stages (Ratke et al.,
2015, 2018).

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 represen-
tation of proteins involved in vesicle formation and move-
ment, and in rosette movement (Table S17, cf. vesicle,
movement category). The identified genes are good candi-
dates for the biotechnological regulation of secondary wall
formation, as the regulation of vesicle and cellulose syn-
thase 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-re-
lated genes, including genes encoding receptor-like
kinases with a lectin domain (Table S17, cf. 'signaling' cat-
egory). 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
<figure><img src="image_6.png" /></figure>

<figure><img src="image_7.png" /><figcaption>**Figure 4.** The difference in neighborhoods of genes encoding proteins of secondary wall xylan synthase complex *PtGT43B* and *PtGT43C* (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 *PtGT43C* (in a) and *PtCesA8-B* (in b) and the orange nodes are unique for *PtGT43B* (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.</figcaption></figure>

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-Teebles *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.

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 *PtCesA8-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 cel-
lulose biosynthetic proteins (for example, *PtCesA4*, *PtCes7-
A*, *PtCes7-B* and *PtGH19A*), but showed differences for tran-
scription 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 tran-
scription 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 biosyn-
thetic genes strictly expressed during secondary wall for-
mation, PtGT43A and PtGT43B (Ratke et al., 2015),
suggesting a secondary wall function for PtCesA8-A (Fig-
ure 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) (Fig-
ure 4b; Table 3). The presence of the AtET01 homolog in
this subnetwork suggests its negative correlation with ethy-
lene signaling; AtET01 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 syn-
thase complexes in this tissue (Zhang et al., 2018).

CONCLUSION

This study provides a census of all CAZymes and expan-
sin-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 rela-
tive size of certain families, suggesting their adaptive regu-
lation. 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), pro-
vided evidence for the expression for 94% of the gene mod-
els, and the expression of 62% of the models in developing
wood tissues, and enabled comparative analyses of expres-
sion. We used the CAZymes expression profiles during
wood development and their relatedness to create corre-
sponding co-expression networks and to identify regulons
composed of CAZymes and other genes. Transcriptional
changes occurring during wood development provided sev-
eral candidates for the biosynthesis of primary and sec-
ondary 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
<table><caption>**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/)</caption><thead><tr><th>Potri. ID</th><th>Potri name</th><th>AT ID</th><th>AT name</th><th>TF class</th><th>CesA7-A<br>Potri.<br>006G181900</th><th>CesA8-A<br>Potri.<br>011G069600</th><th>CesA8-B<br>Potri.<br>004G059600</th><th>GT43B<br>Potri.<br>016G086400</th><th>GT43C<br>Potri.<br>007G047500</th><th>GT47A-1<br>Potri.<br>001G088100</th><th>GATL1-B<br>Potri.<br>002G132900</th><th>GATL1-A<br>Potri.<br>014G040300</th><th>GT47C<br>Potri.<br>009G006500</th><th>CSLA1<br>Potri.<br>008G026400</th><th>GUX1-A<br>Potri.<br>007G107200</th><th>GUX1-B<br>Potri.<br>005G061600</th></tr></thead><tbody><tr><td>001G098800</td><td>MYB10</td><td>1G63910</td><td>MYB103</td><td>MYB</td><td>+</td><td>+</td><td>+</td><td>+</td><td>+</td><td>+</td><td>+</td><td>+</td><td>+</td><td>+</td><td>+</td></tr><tr><td>001G112200</td><td>KNA77</td><td>1G62990</td><td>IXRT1,<br>KNA77</td><td>TALE</td><td>+</td><td>+</td><td>+</td><td>+</td><td>+</td><td>+</td><td>+</td><td>+</td><td>+</td><td>+</td><td>+</td></tr><tr><td>001G268400</td><td></td><td></td><td></td><td></td><td></td><td></td><td></td><td></td><td></td><td></td><td></td><td></td><td></td><td></td><td></td></tr><tr><td>002G119400</td><td></td><td>3G49940</td><td>LBD38</td><td>C3H</td><td>+</td><td>+</td><td>+</td><td>+</td><td>+</td><td>+</td><td>+</td><td>+</td><td>+</td><td>+</td><td>+</td></tr><tr><td>002G257800</td><td></td><td>4G28610</td><td>PHR1</td><td>LBD</td><td>+</td><td>+</td><td>+</td><td>+</td><td>+</td><td>+</td><td>+</td><td>+</td><td>+</td><td>+</td><td>+</td></tr><tr><td>006G253800</td><td></td><td>5G25390</td><td>SHN3</td><td>G2-like</td><td>+</td><td>+</td><td>+</td><td>+</td><td>+</td><td>+</td><td>+</td><td>+</td><td>+</td><td>+</td><td>+</td></tr><tr><td>007G135300</td><td>NAC156</td><td>4G28500</td><td>ANAC073,<br>NAC073,<br>SND2</td><td>ERF</td><td>+</td><td>+</td><td>+</td><td>+</td><td>+</td><td>+</td><td>+</td><td>+</td><td>+</td><td>+</td><td>+</td></tr><tr><td>008G117500</td><td></td><td>1G68870</td><td>HHO2</td><td>G2-like</td><td>+</td><td>+</td><td>+</td><td>+</td><td>+</td><td>+</td><td>+</td><td>+</td><td>+</td><td>+</td><td>+</td></tr><tr><td>009G045900</td><td></td><td>5G60440</td><td>AGL62</td><td>M-type,<br>MADS</td><td>+</td><td>+</td><td>+</td><td>+</td><td>+</td><td>+</td><td>+</td><td>+</td><td>+</td><td>+</td><td>+</td></tr><tr><td>017G016700</td><td>NAC154</td><td>4G28500</td><td>ANAC073,<br>NAC073,<br>SND2</td><td>NAC</td><td>+</td><td>+</td><td>+</td><td>+</td><td>+</td><td>+</td><td>+</td><td>+</td><td>+</td><td>+</td><td>+</td></tr><tr><td>002G031000</td><td>BLH3</td><td>2G16400</td><td>BLH7</td><td>TALE</td><td>+</td><td>+</td><td>+</td><td>+</td><td>+</td><td>+</td><td>+</td><td>+</td><td>+</td><td>+</td><td>+</td></tr><tr><td>002G178700</td><td>WND2B,<br>SND1-B2</td><td>2G46770</td><td>ANAC043,<br>EMB2301,<br>NST1</td><td>NAC</td><td>+</td><td>+</td><td>+</td><td>+</td><td>+</td><td>+</td><td>+</td><td>+</td><td>+</td><td>+</td><td>+</td></tr><tr><td>003G132000</td><td>MYB128</td><td>1G63910</td><td>MYB103</td><td>MYB</td><td>+</td><td>+</td><td>+</td><td>+</td><td>+</td><td>+</td><td>+</td><td>+</td><td>+</td><td>+</td><td>+</td></tr><tr><td>006G138500</td><td></td><td>5G20730</td><td>ARF7.BIP,<br>IAA21,<br>-23-25,<br>MSG1,<br>NPH4,<br>TIF5</td><td>ARF</td><td>+</td><td>+</td><td>+</td><td>+</td><td>+</td><td>+</td><td>+</td><td>+</td><td>+</td><td>+</td><td>+</td></tr><tr><td>008G038900</td><td></td><td>4G32890</td><td>GATAB</td><td>GATA</td><td>+</td><td>+</td><td>+</td><td>+</td><td>+</td><td>+</td><td>+</td><td>+</td><td>+</td><td>+</td><td>+</td></tr><tr><td>001G141100</td><td></td><td>5G46690</td><td>bHLH071</td><td>bHLH</td><td></td><td></td><td></td><td></td><td></td><td></td><td></td><td></td><td></td><td></td><td></td></tr><tr><td>012G055700</td><td></td><td>1G73830</td><td>BEE3</td><td>bHLH</td><td></td><td></td><td></td><td></td><td></td><td></td><td></td><td></td><td></td><td></td><td></td></tr><tr><td>005G221400</td><td></td><td>2G42410</td><td>ZFP11</td><td>C2H2</td><td></td><td></td><td></td><td></td><td></td><td></td><td></td><td></td><td></td><td></td><td></td></tr></tbody></table>

+, positive; -, negative correlation
expression pattern encompassing the cambium and radial
expansion zone. Moreover, the data provide support for the
diversification of functions of the *Populus* secondary cellu-
lose synthase paralogs *PtCesA8-A* and *PtCesA8-B*. The col-
lection 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 data-
base 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 avail-
able 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-informa-
tics/1283-guidelines-for-rna-seq-data-analysis). Briefly, the
quality of the raw sequence data was assessed using
FASTOC (http://www.bioinformatics.babraham.ac.uk/projec
ts/fastqc/). Residual ribosomal RNA (rRNA) contamination
was assessed and filtered using SORTMERNA 2.1 (with set-
tings -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 pro-
vided 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 MIN-
LEN:50; Bolger et al., 2014). After both filtering steps,
FASTOC 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 -quant-
Mode TranscriptomeSAM -outFilterMultimapNmax100 -
outReadsUnmapped Fastx -chimSegmentMin1 -

outSAMtype BAM SortedByCoordinate -outWigType bed-
Graph -alignIntronMax11000; Dobin et al., 2013). The anno-
tations obtained from the *P. trichocarpa* v3.0 GFF file were
flattened to generate 'synthetic' gene models. This syn-
thetic 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 val-
ues. 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 transforma-
tion (VST) in R 3.4.0 (R Core Team, 2015) using the BIOC-
DUCTOR 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 decreas-
ing 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-cen-
tering approach. The resulting data were subjected to hier-
archical 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 avail-
able from our GitHub repository (https://github.com/
UPSCb/UPSCb/tree/master/manuscripts/Kumar2018) and
the corresponding data are available from our FTP reposi-
tory (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 repli-
cates. Arabidopsis thaliana orthologs of the identified
poplar genes were identified by BLAST (https://www.arabid
opsis.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 cus-
tom 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.pop
genie.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-expres-
sion 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 cy-
toscape 3.4.0 (Shannon et al., 2003) for visualization, color-
ing and determining the first neighbors of functionally
known 'Guide Genes' involved in the development of pri-
mary 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 correspond-
ing 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.igi.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 monosaccha-
ride composition by alditol acetates (Englyst and Cum-
mings, 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 previ-
ously (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.

ACKNOWLEDGEMENTS

We thank a talented undergraduate student Tristan
Coustes for help in cell wall analyses.

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 Sys-
tems) and KAW (The Knut and Alice Wallenberg Founda-
tion), 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 analy-
ses. P.I. was responsible for cell wall analyses. EJM con-
ceived and coordinated the project, finalized the
manuscript with contributions from all co-authors, and
agrees to serve as the author responsible for correspon-
dence.

CONFLICT OF INTEREST

The authors declare no conflicts of interest.

SUPPORTING INFORMATION

Additional Supporting Information may be found in the online ver-
sion of this article.

**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 devel-
opmental 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 expres-
sion 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.

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