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# Unveiling lignocellulolytic potential: a genomic exploration of bacterial lineages within the termite gut

João Felipe M. Salgado<fnref n="1" />, Vincent Hervé<fnref n="1" />, Manuel A. G. Vera<fnref n="1" />, Gaku Tokuda<fnref n="2" /> and Andreas Brune<fnref n="1" />*

**Abstract**

**Background** The microbial landscape within termite guts varies across termite families. The gut microbiota of lower termites (LT) is dominated by cellulolytic flagellates that sequester wood particles in their digestive vacuoles, whereas in the flagellate-free higher termites (HT), cellulolytic activity has been attributed to fiber-associated bacteria. However, little is known about the role of individual lineages in fiber digestion, particularly in LT.

**Results** We investigated the lignocellulolytic potential of 2223 metagenome-assembled genomes (MAGs) recovered from the gut metagenomes of 51 termite species. In the flagellate-dependent LT, cellulolytic enzymes are restricted to MAGs of *Bacteroidota* (*Dysgonomonadaceae*, *Tannerellaceae*, *Bacteroidaceae*, *Azobacteroidaceae*) and *Spirochaetota* (*Breznakiellaceae*) and reflect a specialization on cellodextrins, whereas their hemicellulolytic arsenal features activities on xylans and diverse heteropolymers. By contrast, the MAGs derived from flagellate-free HT possess a comprehensive arsenal of exo- and endoglucanases that resembles that of termite gut flagellates, underlining that *Fibrobacterota* and *Spirochaetota* occupy the cellulolytic niche that became vacant after the loss of the flagellates. Furthermore, we detected directly or indirectly oxygen-dependent enzymes that oxidize cellulose or modify lignin in MAGs of *Pseudomonadota* (*Burkholderiales*, *Pseudomonadales*) and *Actinomycetota* (*Actinomycetales*, *Mycobacteriales*), representing lineages located at the hindgut wall.

**Conclusions** The results of this study refine our concept of symbiotic digestion of lignocellulose in termite guts, emphasizing the differential roles of specific bacterial lineages in both flagellate-dependent and flagellate-independent breakdown of cellulose and hemicelluloses, as well as a so far unappreciated role of oxygen in the depolymerization of plant fiber and lignin in the microoxic periphery during gut passage in HT.

**Keywords** CAZymes, Lignocellulose degradation, Lignin, Functional genomics, Cellulase, Termite microbiota

**Background**

Termites are eusocial insects that effectively digest wood and other lignocellulosic plant matter [1]. Their degradation mechanism involves a dual system. The termite comminutes the wood particles and degrades amorphous regions of the cellulose with endoglucanases and β-glucosidases secreted in the salivary glands or the midgut [2, 3], and symbiotic microorganisms in the dilated hindgut subsequently degrade the wood particles that pass from the midgut into the hindgut largely intact [4]. The contributions of microbial

<footnote>*Correspondence:</footnote>

<footnote>Andreas Brune
brune@mpi-marburg.mpg.de</footnote>

<footnote n="1">¹ RG Insect Microbiology and Symbiosis, Max Planck Institute for Terrestrial Microbiology, 35043 Marburg, Germany</footnote>

<footnote n="2">² Tropical Biosphere Research Center, Center of Molecular Biosciences, University of the Ryukyus, Nishihara, Okinawa 903-0213, Japan</footnote>
symbionts to fiber digestion are essential as the host lacks both exoglucanases acting on crystalline cellulose and any hemicellulolytic activities.

The microbial community in the hindgut of termites is highly specialized and differs between lineages [5]. Flagellates dominate the microbiota of most termite families (collectively referred to as lower termites, LT), where they sequester wood particles from the hindgut fluid into digestive vacuoles containing both endoglucanases and exoglucanases [55], suggesting that bacteria are of little relevance in the digestion of plant fiber for these termites [5]. In the family Termitidae (higher termites, HT), the loss of cellulolytic flagellates is compensated by the cellulolytic activity of their bacterial microbiota [6, 7]. In the subfamily Macrotermitinae, which cultivate basidiomycete fungi of the genus *Termitomyces* that degrade (hemi)cellulose and depolymerize lignin [8], the need for degradation during gut passage is alleviated by the pretreatment of lignocellulose in fungal gardens [9].

Therefore, the breakdown of recalcitrant fibers in the hindgut of most HT is an entirely bacterial process [2]. Metagenomic studies revealed the presence of carbohydrate-active enzymes (CAZymes) that are often closely related to those of cellulolytic clades from the rumen of cattle or other environments [10–12]. However, these studies covered only a few species of higher termites, and from the arsenal of glycoside hydrolases (GHs), only a few cellulases (GH5) of unidentified hindgut bacteria and hemicellulases (GH10 and GH11) of *Treponematales* have been characterized [12–14].

Apart from a recent study, which analyzed the CAZymes in a wide range of metagenomes from many termite families and documented considerable differences in the CAZyme repertoire of LT and HT [15], little is known about the fiber-digesting potential of specific bacterial lineages of the gut microbiota in LT. Most importantly, only few of the CAZymes identified in the metagenomic datasets have been pinpointed to specific bacterial lineages using metagenome-assembled genomes (MAGs) [10, 16, 18]. Moreover, most previous analyses focused on GHs but did not discuss other CAZyme families acting on lignocellulose, such as carbohydrate-binding modules (CBM) and auxiliary activities (AA). CBMs are enzyme domains involved in the attachment of GHs to their respective substrate, whereas some AAs participate in redox reactions that depolymerize (hemi)celluloses and lignin and are directly or indirectly dependent on molecular oxygen as co-substrate [17]. Certain carbohydrate esterases (CE) remove methyl and acetyl groups from the polysaccharide backbone of hemicelluloses and, together with polysaccharide lyases (PL), also contribute to pectin degradation. Glycosyl transferases (GT) are

mostly involved in the biosynthesis of glycoproteins and glycolipids [18–20].

Here, we present a comprehensive genome-centric analysis of the CAZymes encoded by more than 2000 bacterial MAGs reconstructed from gut metagenomes of 51 termite species [21, 22]. Using a vast genomic library, covering nearly all termite families, we identify the distribution of CAZymes among all major bacterial lineages previously detected in termite guts. We focus on GHs, AAs, and CBMs with lignocellulolytic activity among MAGs from individual bacterial lineages from different groups of termites, for the first time including also oxygen-dependent activities.

## Methods

### Metagenome-assembled genomes and phylogenomics

MAGs were generated in previous metagenomic studies from our group [21, 22]. Metadata associated with the corresponding metagenomes can be found in Supplementary Table S1. The quality of the reconstructed genomes was estimated with CheckM v1.1.6 [23]. MAGs that were at least 50% complete and less than 10% contaminated were retained for analyses. Relative abundances were calculated as previously described [22]. MAGs were classified using the Genome Taxonomy Database (GTDB) with the GTDB-Tk v2 toolkit [24], based on an alignment of 120 ubiquitous marker genes retrieved from the MAGs, and phylogenies were reconstructed using IQ-TREE [25]. To validate genome classifications, taxonomies from Pplacer v1.1.17 [26] and CheckM were taken into consideration. Metadata associated with each MAG can be found in Supplementary Table S2. The termite phylogeny is based on previous work [15, 27, 28], using a combination of rRNA genes and mitochondrial genomes. All maximum-likelihood phylogenetic trees were curated using the packages ape [29] and PAML [30]. Genomic data were mapped to phylogenies using iTOL [31] and R package *ggtree* [32].

### Gene prediction and annotation

Genomes were subjected to gene prediction using Prodigal [33], with 4,480,144 genes predicted. Coding densities were also estimated by the software, by averaging the total CDS bases as a function of the total genome bases and expressed in percentages, and can be found in Table S2. Protein domain annotations were added using the TIGRFAM [34], Pfam [35], SUPERFAMILY [36], Panther [37], and CDD [38] databases in the INTERPROSCAN [39] classifier, with default parameters. CAZymes were identified using Hidden Markov Model (HMM) searches (e-value < 1e-15, score > 100, and coverage > 0.35) against the dbCAN3 and dbCAN-sub metaservers [40], using the function *run_dbcan*. The classification was
confirmed with protein domain annotations and DIA-
MOND [41] searches (e-value < 1e-15, id > 50%) against
the CAZy database [42]. Signal peptides were identified
with SignalP V.6 [43] and Phobius [44].

Multivariate analyses and cluster visualization

A dataset including all high-quality CAZyme genes
with secretion signals was subjected to uniform mani-
fold approximation and projection (UMAP), using the
R package *umap* [45]. The k-nearest neighbors (KNN)
algorithm was used with parameters for 15 neighbors and
500 epochs for training. For the input, the raw-count data
were square-root transformed and Wisconsin double
standardized with the function *decostand*, and a Bray-
Curtis dissimilarity matrix was calculated using the func-
tion *vegdist*; both functions are from the R package *vegan* [46].
The analysis included host family and subfamily,
termite diet, gut compartment, and bacterial phylum as
metadata. The contribution of these variables to CAZyme
composition was further estimated with the functions
ADONIS and ANOSIM, also based on Bray-Curtis dis-
similarities, each with 1000 permutations and boot-
strapped 100× for best model fit (max stress = 0.12) in
the *vegan* package. Principal component analyses (PCA)
were used as implemented in the function *prcomp* of the
R package *stats* [47]. Explanatory PCs were employed in
rarefactions for the relationship between CAZyme family
distributions in relation to either the per-MAG number
of genes predicted or metagenomic relative abundance.

CAZyme family selection and statistical analyses

Outliers were removed, and homogeneity of variance
and normality was checked using Levene's and Shapiro
Test from the R packages *car* [48] and *stats*. Since neither
assumption was applicable, a nonparametric Kruskal–
Wallis test was used on the entire pre-selection CAZyme
dataset to determine significantly different CAZyme
families across the 15 bacterial phyla, with the function
*kruskal.test* and with post hoc Mann–Whitney *U* tests
in the function *wilcox_test* of the package *rstatix* [49],
followed by Bonferroni multiple testing correction at a
threshold of *p* < 0.0001, using the *pzeadjust* function of the
*stats* package. A total of 146 CAZyme families tested pos-
itive for association with the phyla, of which those with
lignocellulolytic activities (38 CAZyme families contain-
ing 8 CBMs) were selected for downstream comparison.

The classification of individual CAZymes into the
major lignocellulolytic categories (cellulase, hemicel-
lulase, ligninase, chitinase, and pectinase) was based
on the gene-wise presence and complementary activi-
ties using the substrate predictions of DBCAN-sub (see
Table S3). Enzymes with only endo-β-1,4-glucanase
activities or other classically cellulolytic activities, such

as cellobiohydrolases, were classified as cellulases,
whereas enzymes with endo-β-1,4-glucanase activity
and additional xylanolytic or xyloglucan-specific activi-
ties were classified as hemicellulases. Enzymes from a
given CAZyme family with hemicellulolytic activities
were considered hemicellulases even if they comprised
endo-β-1,4-glucanase activity. Enzymes from family GH3
were not included in this categorization given the com-
plexity of their predicted substrates. CBMs that were not
connected to CAZymes assigned to specific GH families
were removed from the dataset. CAZymes from fami-
lies with peroxidase, aryl alcohol oxidase, and/or lac-
case activities were classified as ligninases. Pectate lyases
and pectin methylesterases were classified as pectinases.
Enzymes with acetylglucosaminosidase, chitin deacety-
lase, and chitinase activities were classified as chitinases.

To confirm bacterial phylogeny as a covariate, we
employed a local indicator of phylogenetic associa-
tion (lipaMoran) analysis coupled with an Abouheif
test. CAZyme families containing (hemi)cellulases and
lignin-modification enzymes were used as traits, and the
phylogenetic signal was considered at *p*-values < 0.05.
Correlograms were plotted with the R package *phylosig-
nal* [50]. Since there was no considerable coding density
variation between MAGs, gene densities (GD) were esti-
mated by normalizing the numbers of genes in the ligno-
cellulolytic CAZyme classifications by per-MAG number
of predicted genes. Specific Wilcoxon's signed-rank tests
were used to assess the association of GD in HT and LT,
and the effect sizes were estimated using the function
*wilcox_effsize* of the package *rstatix*. Correlation between
continuous variables was tested with Spearman correla-
tion rank, as implemented by the function *cor.test* in the
R package *stats*.

Results

Representation of bacterial MAGs and *phyla* in termite microbiomes

The initial dataset comprised 2223 bacterial MAGs that
were recovered from 51 termite species, spanning seven
families of lower termites (LT) and seven subfamilies
of higher termites (HT) (Fig. 1). Archaeal MAGs were
excluded because preliminary analyses revealed negli-
gible numbers of CAZymes. The MAGs represented 22
bacterial phyla, with the majority classified among *Bacil-
lota* (24.1%), *Bacteroidota* (15.6%), *Spirochaetota* (14.2%),
and *Pseudomonadota* (13.4%).

The MAGs from different bacterial phyla differed
strongly in relative abundance, ranging between 0.004
and 19.8% (average 0.31%) of the reads in the corre-
sponding metagenomes (Fig. 1). Certain phyla (e.g.,
*Elusimicrobiota* and *Verrucomicrobiota*) were more prev-
alent among lower termites (LT), whereas others were
<figure><img src="image_2.png" /><figcaption>**Fig. 1** Distribution of MAGs from major bacterial phyla among metagenomes from different host groups. The ordinate shows the number of MAGs in each phylum as a heatmap that is based on the total number of MAGs (2,223) in the dataset. The abscissa shows the number of metagenomes from which the MAGs were recovered, summarized for each host group, as a heatmap based on the total number of metagenomes (51). Relative abundance values indicate the proportion of reads in a metagenome that mapped against the corresponding MAGs, expressed as averages per termite family (lower termites) or subfamily (higher termites, Termitidae). The termite species contained in each host group, the relative abundances of individual MAGs, and their classification down to genus level are given in Tables S1 and S2</figcaption></figure>

more prevalent (*Pseudomonadota*) in or even restricted to (*Fibrobacterota*) higher termites (HT). Many phyla were represented only in certain (sub)families of LT and HT but virtually absent in others (e.g., *Actinobacteriota*, *Bacteroidota*, *Desulfobacterota*, *Spirochaetota*, and *Patescibacteria*). The bacterial phyla *Chlamydiota*, *Cyanobacteriota*, *Myxococcota*, *Cloacimonadota*, *Chloroflexota*, *Fusobacteriota*, and *Deferribacteria* presented less than 5 MAGs and were removed from subsequent analyses.

The remaining 2204 MAGs from 15 bacterial phyla were analyzed for CAZyme distribution. From the full dataset (Table S5), we selected 38 CAZyme families with predicted secretion signals that are involved in lignocellulose degradation (Table S3) and were differentially distributed across phyla ($p \le 0.001$) (Fig. 2). They comprised 18 families with cellulolytic, 13 families with hemicellulolytic, and 7 families with lignin-modifying activities

(Table S3), possessing additionally 8 CBM domain families. CAZyme families that are not directly involved in lignocellulolytic processes, such as glycosyltransferases (GT) and polysaccharide lyases (PL), were excluded from the analysis. Carbohydrate esterases (CE) of subfamily CE4 were represented in almost all MAGs (Table S5), but since their predicted substrates did not include acetylxylan, they were also excluded.

There were clear patterns in the distribution of the selected CAZymes among bacterial lineages (Fig. 2). Both cellulolytic and hemicellulolytic activities were prominent among the MAGs of *Bacteroidota*, especially *Sphingobacteriaceae*, *Chitinophagaceae*, and most families of *Bacteroidales*. MAGs of the *Fibrobacterota* possessed a high abundance of cellulolytic enzymes, with specific sets of CAZyme families that were not observed in other phyla. Large numbers of (hemi)cellulases were present also in MAGs of the
<figure><img src="image_3.png" /><figcaption>**Fig. 2** The distribution of carbohydrate-active enzymes with lignocellulolytic activities (cellulase, hemicellulase, auxiliary) and cellulose-binding modules (CBM) among the MAGs of major bacterial families recovered from termite gut metagenomes. The heatmap indicates the mean gene abundance in selected CAZyme families for the MAGs from the respective bacterial family. CAZYme families were clustered using a hierarchical clustering algorithm; values were scaled from 0 to 1 to facilitate visualization. An interactive spreadsheet with details for all MAGs is given in Table S4; the complete dataset is given in Table S5</figcaption></figure>

*Bacillota* and *Actinobacteriota*. Among *Bacillota*, MAGs with considerable numbers of cellulases were found mostly among *Clostridia* (e.g., *Lachnospiraceae*, *Acutalibacteraceae*, and *Clostridiceae*), whereas the only prominent family with cellulases among *Bacilli*

was *Paenibacillaceae*. Clear differences were apparent also in the distribution of auxiliary activities and CBMs (see below).
## Factors driving bacterial CAZyme distribution

Based on the obvious patterns in CAZyme distribution among MAGs from different bacterial lineages, we went back to the entire pre-selection dataset of CAZymes (256,212 genes across 534 CAZyme families; Table S5) to test whether the distribution patterns of CAZymes in the termite gut reflect bacterial taxonomy or host factors. Dimensionality reduction using all CAZyme families revealed that the bacterial phylum is an almost exclusive driver of CAZyme distribution (Fig. 3A), followed to a lesser extent by the (sub)family of its termite host (Fig. 3B). A separation of the MAGs from LT and HT reveals similar distribution patterns (Fig. S1A-B). Other factors, such as host diet or gut compartment, did not affect the ordination (Fig. S1C-D). A fitted phylogenetic signal model confirmed the association of bacterial taxonomy to the 38 selected CAZyme families. There was phylogenetic signal in distribution of cellulases in the phyla *Bacillota*, *Actinomycetota*, *Bacteroidota*, *Fibrobacterota*, and *Planctomycetota* (Figs. S2 and S3). The phylum *Spirochaetota* presented a phylogenetic signal for the distribution of hemicellulases. *Actinomycetota*, *Pseudomonadota*, and *Acidobacteriota* were the only phyla with a significant phylogenetic signal for the ligninases.

## Division of fibrolytic niches among bacterial clades

Using the ratio of (hemi)cellulase genes in the selected 38 CAZyme families to genes per genome (gene density, GD) to allow for abundance comparison between clades, we found that MAGs with the highest GD of cellulases were *Fibrobacterota*, followed by *Bacteroidota*, *Bacillota*, and *Spirochaetota* (Fig. 4A). The bacterial phyla with the highest hemicellulolytic potential were *Bacteroidota*, *Bacillota*, and *Spirochaetota*, followed by *Fibrobacterota* and *Verrucomicrobiota* (Fig. 4B).

Among *Fibrobacterota*, a cluster of cellulase-containing CAZymes was highly abundant in MAGs of all families (*Fibrobacteraceae*, *Chitinivibrionaceae*, and *Chitinispirillaceae*). It is composed of GH9, GH5 (subfamilies 5, 2, and 39), GH8, and GH94 (Figs. 2 and 5A). The GH9s from our bacterial dataset comprise mainly endo-β-1,4-glucanases but only very few exo-β-1,4-glucanases, while GH8 and the specified GH5 subfamilies contain only endo-β-1,4-glucanases (Table S3). GH94 is composed mainly of enzymes with cellodextrin phosphorylase (CDP) and cellobiose phosphorylase (CBP) activity (Table S3). Although the enzymes characterized to date are generally cytoplasmic, one-third of the GH94 enzymes in our dataset have a signal peptide (Table S6). Also, MAGs of the *Oscillospiraceae*, *Lachnospiraceae*,

<figure><img src="image_4.png" /><figcaption>**Fig. 3** Ordination analysis of CAZyme composition in 2,204 MAGs of termite-associated bacteria. The UMAP analysis (stress = 0.12) used the entire dataset of CAZymes (256,212 genes across 534 CAZyme families; see Table S5) to visualize differences in the general content of CAZymes among phyla. The biplot displays MAGs color-coded according to their respective phyla (A) and according to the termite (sub)families from which they were recovered (B). The theoretical distances were calculated based on the KNN score, and the statistics are based on ANOSIM (bacterial phyla R=0.34; termite (sub)families R=0.04; *p* < 0.001) and ADONIS (bacterial phyla R²=0.24; termite (sub)families R² = 0.03; *p* < 0.001) tests</figcaption></figure>
<figure><img src="image_5.png" /><figcaption>**Fig. 4** Gene density (GD) of cellulases (A) and hemicellulases (B) in the termite-associated MAGs of different bacterial phyla, and the relationships between the respective GDs and the relative abundance (RA) of the MAGs in the metagenomes of lower termites and higher termites (C, D). The number of (hemi)cellulolytic genes of a given MAG is normalized by its total number of predicted genes and expressed as percentages. Statistical tests were performed both globally and pairwise, against the mean values (dashed line). Significance values: ns = non-significant; \* ≤ 0.05; \*\* ≤ 0.01; \*\*\* ≤ 0.001; \*\*\*\* ≤ 0.0001. Linear regressions are shown for all phyla but a positive correlation (Spearman ρ > 0.1 and p < 0.05) was present only for LT-associated MAGs of *Bacteroidota* and HT-associated MAGs of *Fibrobacterota*, *Bacteroidota*, *Spirochaetota*, *Bacillota* and *Actinomycetota*</figcaption></figure>

and *Clostridiaceae* (*Bacillota*) possessed CDPs and CBPs, albeit in lower abundance (Figs. 2 and 5A).

The most abundant GH family in all lineages of *Bacteroidota* was GH3 (1,139 genes), corresponding to 3.3 genes per genome (GPG). Members of GH3 have β-glucosidase activity with wide substrate specificity, precluding classification as cellulases or hemicellulases (Table S3). Additionally, the orders *Bacteroidales*, *Chitinophagales*, and *Sphingobacteriales* displayed consistent abundances of GH16 (subfamilies 4, 5, and 3) and GH30 (subfamilies 1, 6, and 3; Figs. 5A and 6A). These subfamilies contained lichenases/laminarinases with endo-β-1,3- and endo-β-1,4-glucanase activity (Table S3) [98].

Among *Spirochaetota*, MAGs of *Breznakiellaceae*, *Spirochaetaceae*, and *Treponemataceae* had high abundances of GH11, GH10, GH5_22, GH5_4, and GH5_8, which contribute strongly to their divergence in hemicellulolytic functions from other phyla (Fig. 6B). Enzymes from these families have a variety of substrates, acting mainly on xyloglucans and xylomannans (Table S3). By contrast, MAGs of *Bacteroidota* showed a high abundance of GH2 (1,734 genes, 5 GPG on average). Enzymes in this family have β-galactosidase and α-arabinosidase activity (Table S3). Additionally, the MAGs from many

families in the orders *Bacteroidales*, *Sphingobacteriales*, and *Chitinophagales* had GH43 genes (Figs. 2 and 5B), which encode endo-xylanase and α-arabinofuranosidase activities (Table S3). The high abundance of GH2 and GH43 genes drives the hemicellulase composition of the *Bacteroidota* MAGs (Fig. 6B), accounting for most of the observed variation. High numbers of GH2 and GH43 genes were present also among MAGs of *Verrucomicrobiota* (*Opitutaceae* and UBA953) and *Bacillota* (*Oscillospiraceae*, *Lachnospiraceae*, *Peptostreptococcaceae*, *Ruminococcaceae*) (Figs. 2 and 5B). However, *Bacillota* MAGs displayed even higher proportions of GH26, with activity on xylomannan (Fig. 5B, Table S3). While none of the CE associated with our dataset were annotated as acetylxylan esterases, we found pectin lyases and CE involved in pectin degradation (pectin acetyl esterases and pectin methyl esterases) abundantly represented in many MAGs (Tables S3, S5), particularly from the phyla *Bacteroidota* and *Planctomycetota* (Figure S6).

## Differences in (hemi)cellulolytic lineages between lower and higher termites

MAGs of *Fibrobacterota* were recovered exclusively from HT, and MAGs of *Spirochaetota* MAGs from HT
<figure><img src="image_6.png" /><figcaption>**Fig. 5** Abundance and proportion of selected CAZymes from GH families with cellulolytic (A) and hemicellulolytic (B) functions in the MAGs of selected bacterial families with a high fibrolytic potential. Each bar shows the total number of CAZymes classified as cellulase or hemicellulase that are encoded by a particular MAG (gray and black color indicates the origin from lower or higher termites) and the proportion of genes in the respective GH families (visualized with different colors). Only MAGs that encode more than one cellulase or hemicellulase were included. The families GH5 and GH30 appear twice but refer to different subfamilies classified as cellulases or hemicellulases (see Table S3 for details)</figcaption></figure>

had a higher gene density of cellulases than those from LT ($p < 0.001$ and $r=0.6$; Fig. 4A). There was a clear correlation between (hemi)cellulase density and metagenomic relative abundances in MAGs of *Fibrobacterota* and *Spirochaetota* from HT (both with $p < 0.001$ and $\rho=0.5$; Fig. 4C–D). Both *Fibrobacterota* (*Fibrobacteraceae*, *Chitinivibrionaceae*, and *Chitinispirillaceae*) and *Spirochaetota* (*Breznakiellaceae*) MAGs from HT had high proportions of GH5, GH8, and GH9 (Fig. 5A). MAGs of *Bacteroidota* and *Bacillota* differed only slightly between LT and HT (both with $p < 0.001$ and effect size

$r < 0.1$; Fig. 3). However, LT-derived genomes of *Dysgonomonadaceae*, *Azobacteroidaceae*, *Bacteroidaceae*, *Tannerellaceae* (*Bacteroidota*), and *Breznakiellaceae* (*Spirochaetota*) had a higher proportion of GH30, GH16, and GH116, which act on cellodextrins (Fig. 5A).

The situation was different in the case of hemicellulases. Here, the GDs were higher in MAGs of *Spirochaetota* and *Bacillota* derived from HT (both with $p < 0.001$ and approx. $r=0.3$; Fig. 4B), with large proportions of GH5, GH10, and GH11 (Fig. 5B). By contrast, MAGs of *Bacteroidota* from LT had higher densities
<figure><img src="image_7.png" /><figcaption>**Fig. 6** Principal component analysis (PCA) of the abundance of cellulases (A) and hemicellulases (B) in the MAGs from selected bacterial families with a high fibrolytic potential (see Fig. 5). The biplots show MAGs color-coded either according to bacterial phylum (left) or according to their respective host group (lower or higher termite; right)</figcaption></figure>

of hemicellulases than those from HT (*p* < 0.001 and *r* = 0.2). LT-derived genomes of *Dysgonomonadaceae*, *Azobacteroidaceae*, *Bacteroidaceae*, *Tannerellaceae* (*Bacteroidota*), and *Breznakiellaceae* (*Spirochaetota*) had a higher proportion of GH2 and GH43, which comprise CAZymes with endoxylanase, arabinofuranosidase and β-galactosidase activities (Fig. 5B). LT-derived *Bacteroidota* MAGs with high GDs of (hemi) cellulases were abundant in the metagenomes, although the correlation between GD and relative abundance was not strong (*p* < 0.1, Spearman ρ < 0.1; Fig. 4C–D).

The ordination analysis reveals that the prevalence of CAZymes with different activities was driven not only by differences in taxonomy but also by their colonization patterns of LT and HT (Fig. 6).

**Carbohydrate-binding modules in CAZymes from (hemi) cellulolytic clades**

Carbohydrate-binding modules (CBMs) are domains responsible for the adherence of certain CAZymes to their ligand, assuring that secreted enzymes do not diffuse away from their substrates. Among MAGs of the
phylum *Fibrobacterota*, CBM4 and CBM6 were highly abundant in CAZymes from GH8, GH9, and GH5 (62 genes, average 2 GPG; 23 genes, average 0.7 GPG, respectively). Among MAGs of *Bacteroidota*, CBM6 and CBM62 were represented particularly among CAZymes from GH3 and GH2 from *Prolixibacteraceae*, *Bacteroidaceae*, *Tannerellaceae*, *Dysgonomonadaceae*, and *Azobacteroidaceae* (Fig. 2). MAGs of *Bacillota* showed a low content of CBM9, CBM22, and CBM35 (on average, max. 0.4 GPG) in CAZymes from GH2, GH43, and the GH5 subfamilies with hemicellulase activity. Cohesins, which are characteristic of the cellulosomes of *Bacillota*, were generally present only in *Oscillospiraceae* (average 1.12 GPG) but rare in *Ruminococcaceae* (average 0.3 GPG) (Table S4).

Auxiliary activities involved in fiber digestion and lignin
modification

CAZyme families classified as “Auxiliary Activities” (AA)
comprise different classes of oxidative enzymes that,
among other substrates, can act on lignin or (hemi)cel-
lulose (Table S3). They were less abundant than the
GHs and restricted to MAGs from selected lineages
(Figs. 2 and 7). Lytic cellulose/chitin monooxygenases
(LPMO; AA10, 92 genes) and cellooligosaccharide dehy-
drogenases (CDH; AA7, 1068 genes) were most abundant

among MAGs of *Pseudomonadota* (0.5 GPG on average) and *Actinobacteriota* (0.7 GPG), particularly in the orders *Burkholderiales*, *Propionibacteriales*, *Actinomycetales*, and *Microccocales* (Fig. 7).

CAZymes acting on lignin or its derivatives were also present among MAGs from *Pseudomonadota* and *Actinomycetota* (Fig. 7). They comprised phenol-oxidizing laccases (POL; AA1), lignin-modifying peroxidases (LMP; AA2), and aryl alcohol oxidases (AAO; AA3_2). CAZymes of these families were also found in MAGs from several orders of *Bacteroidota* (mainly LMP) and *Bacillota* (mainly POL) (Fig. 7). Notably, most MAGs that encode lignin-modifying activities were derived from the guts of higher termites (p < 0.05 and r=0.25).

Discussion

Two events in the evolutionary history of termites
involved fundamental changes in the strategy of symbi-
otic digestion: The first event was the acquisition of cel-
lulolytic flagellates by the common ancestor of termites
and wood roaches (Cryptocercidae), which are essential
for the degradation of plant fibers in lower termites (LT).
The second event was the loss of these flagellates in a
common ancestor of higher termites (HT), which led to
functional changes that shifted lignocellulose digestion to
the prokaryotic gut microbiota [5, 51].

<figure><img src="image_8.png" /><figcaption>**Fig. 7** Distribution of gene densities in CAZymes with lignin-modifying activities in MAGs from different bacterial phyla. The insets break down selected phyla to the order level and show the proportion in percentages of genes from different enzyme families: POL (phenol-oxidizing laccases, AA1), LMP (lignin-modifying peroxidases, AA2), and AAO (aryl alcohol oxidases, AA3_2)</figcaption></figure>
The finding that most of the cellulolytic activity in the hindgut of *Nasutitermes* spp. (Nasutitermitinae) is associated with wood particles implicated in fiber-associated bacterial lineages in the degradation of (hemi)celluloses [6, 52]. Wood particles are colonized predominantly by members of the phyla *Fibrobacterota* (*Fibrobacterales* and *Chitinivibrionales*) and *Spirochaetota* (mostly *Breznakiellaceae*) [14, 52]. Our results corroborate previous metagenomic and metatranscriptomic analyses of *Nasutitermes*, *Amitermes*, *Cortaritermes*, and *Microcerotermes* species [10, 14, 16, 53], extending the cellulolytic potential of *Fibrobacteraceae* and *Chitinivibrionaceae* and the hemicellulolytic potential of *Breznakiellaceae* to MAGs from almost all HT investigated, and documenting a cellulolytic potential also in MAGs of the family *Chitinispirillaceae*. The consistent presence of endo-glucanases and β-glucosidases (GH8, GH9, and certain subfamilies of GH5) — accounting for the highest cellulase gene densities of all phyla — indicates that all lineages of *Fibrobacterota* in the hindgut of HT are specialized in cellulose degradation. Conversely, high gene densities of endo-xylanases, β-xylosidases, and endo-mannanases (GH11, GH10, and certain subfamilies of GH5) identify MAGs of *Breznakiellaceae* (*Spirochaetota*) as important hemicellulose degraders in the hindgut of HT. This expands previous evidence for high expression of GH11 and GH5 transcripts in *Nasutitermes* and *Cortaritermes* sp. [10, 14], suggesting a functional specialization among the lineages of *Breznakiellaceae* associated with HT.

Notably, the (hemi)cellulolytic functions in the genomes of fiber-associated bacteria from HT resemble the activities revealed by metatranscriptomic analyses of the cellulolytic flagellates in LT [54, 55]. While the cellulolytic functions of the flagellates comprise exo-glucanases (GH7) and endo-glucanases (GH45) that were not prominent in the bacterial MAGs, the hemicellulolytic functions of the flagellates, i.e., arabnosidases (GH43), endomannases (GH26), and endo-β-1,4-xylanases (GH8, GH10 and GH11), were the same as those encoded by the fiber-associated bacteria in HT. Apparently, the loss of the (hemi)cellulolytic functions of the gut flagellates selected for bacterial lineages with different cellulolytic strategies but a similar arsenal of hemicellulolytic enzymes.

Obviously, the prediction of substrate specificity has clear limitations, and not all CAZymes in our dataset could be attributed to a specific function. This may obfuscate the results of quantitative analyses, such as phylogenetic signal or functional variability, which turned out to be remarkably similar in all instances for flagellases and hemicellulases. Furthermore, the classification of CAZymes involves simplifications, especially when enzymes that act exclusively on cellulose or hemicellulose

fall into the same subfamily, or when the enzyme in question is multifunctional. Nonetheless, the fact that our qualitative analyses revealed clear differences in the subfamilies of CAZymes between particular bacterial lineages corroborates that the classifications used in our study were productive and appropriate for this dataset.

The large number of CBMs in cellulases of *Fibrobacterota* MAGs and the presence of hemicellulose-binding CBM9 in *Breznakiellaceae* MAGs are consistent with the localization of the cellulolytic and hemicellulolytic activities with the particulate fraction both in the termite gut [14, 52] and in the rumen [56]. The organization and mechanisms by which these activities are secreted are, nevertheless, still elusive in both phyla. Moreover, CBMs were encoded also by MAGs from other phyla, such as *Verrucomicrobiota*, *Bacteroidota*, and *Actinomycetota* (Fig. 2). By contrast, CBMs were rare among the diverse lineages of Clostridia with (hemi)cellulolytic potential.

The dominant cellulases in the MAGs of most clostridial lineages were cellobiose and cellodextrin phosphorylases (CBP and CDP; GH94). The CBPs and CDPs characterized to date are cytosolic enzymes that help anaerobic bacteria conserve metabolic energy by an intracellular cleavage of cellodextrins or cellobiose into glucose and glucose 1-phosphate [57, 58]. It was therefore quite unexpected that about a third of GH94 enzymes encoded by termite-associated MAGs carried a signal peptide. So far, extracellular GH94 with CBP/CDP activities have been reported only in a single proteomic study of the clostridial *Caldicellulosiruptor bescii* [59]. The presence of potentially secreted CBP/CDPs in MAGs from both LT and HT suggests that these bacteria utilize cellobiose and dextrins produced by other, cellulolytic microbiota and may explain the low number of CBMs and virtual lack of cellulosomes among the clostridial lineages.

An analysis of lignocellulose digestion by solid-state NMR has revealed significant differences in the fate of polysaccharides between wood-feeding LT and HT [60]. While both *Cryptotermes* (LT) and *Nasutitermes* (HT) efficiently degrade cellulose, the latter significantly outperforms the former in the breakdown of hemicelluloses. Apparently, the breakdown of hemicelluloses, whose structural complexity requires the concerted action of numerous enzymes [61], is accomplished more efficiently by the consortium of bacterial species that colonize the wood particles in the hindgut of HT than by the hemicellulases produced by the gut flagellates of LT [62–64]. In LT, the wood particles are sequestered into the digestive vacuoles of the flagellates as soon as they pass the enteric valve [65], precluding a primary role of gut bacteria in the breakdown of (hemi)cellulose. However, the consistent presence of β-glucosidase and cellodextrinases with
CBMs in the MAGs of *Bacteroidales* associated with LT (*Bacteroidaceae*, *Dysgonomonadaceae*, *Tannerellaceae*, and *Azobacteroidaceae*) suggests that these lineages are involved in the breakdown of cellobiose and cellodextrins, which either stem from the partial degradation of amorphous cellulose by the endogenous cellulases in the midgut [66] or represent products of this incomplete digestion of cellulose by the flagellates. Such task partitioning between bacteria and flagellates had been postulated already to explain the considerable differences in the repertoire of bacterial CAZymes between LT and HT [15].

The utilization of partially digested hemicelluloses by *Bacteroidales* in LT is consistent with the compositional differences in hemicellulolytic activities to HT-associated *Breznakiellaceae* (*Spirochaetota*) and the presence of polysaccharide utilization loci (PUL) that encode both carbohydrate capture and uptake systems and CAZymes for the degradation of cellulose, xylans, and mannans in all lineages of *Bacteroidota* from termite guts [15, 67]. Notably, many of the MAGs from LT represent bacterial lineages associated with gut flagellates [22]. While the ectosymbiotic members of *Azobacteroidaceae* (*Candidatus* Symbiothrix dinenymphae), *Dysgonomonadaceae*, and *Bacteroidaceae* show the same (hemi)cellulolytic potential as their free-living relatives [68–70], the situation is different for the intracellular symbionts (*Candidatus* Azobacteroides pseudotrichonymphae) present in almost all Rhinotermidae [71, 72], whose genomes lack (hemi)cellulolytic potential [73] (Table S4).

Since the polyphenolic component of lignocellulose provides a major obstacle to the enzymatic attack of cellulose and hemicelluloses, the fate of lignin during termite gut passage has been disputed for decades [74]. The mechanistic challenges of lignin degradation are rooted in the stability of the inter-monomeric linkages, which cannot be hydrolyzed but require oxidative attack by (per)oxidases [75]. Meanwhile, there is compelling evidence that lignin is modified during gut passage in all termites investigated [76–79]. Nonetheless, there are fundamental differences between LT and HT. While LT disrupts the covalent bonds between hemicelluloses and lignin, leaving the latter largely intact, HT efficiently degrade the lignin polymer, as evidenced by the depletion of major inter-unit linkages and methoxyls [78, 79].

The capacity for the breakdown of lignin is found in many soil bacteria [81], which possess the same types of extracellular (per)oxidases that are present in lignin-degrading fungi [75, 81, 82]. Many of these lineages were represented among our termite MAGs, particularly within *Actinomycetota* (*Micrococcales*, *Actinomycetales*, and *Mycobacteriales*) and *Pseudomonadota* (*Burkholderiales* and *Pseudomonadales*). The MAGs of each lineage

encoded a diverse set of laccases, lignin peroxidases, and aryl-alcohol oxidases. Laccases (phenol oxidases) are the most important agents of bacterial lignin degradation that not only oxidize phenolic substrates but — in the presence of lignin degradation products as mediator — also cleave the inter-unit bonds of the lignin polymer [83]. The same bonds are cleaved also by lignin peroxidases, which require hydrogen peroxide produced by aryl-alcohol oxidases as co-substrate [75].

It is noteworthy that almost all MAGs with lignin-modifying capacity were recovered from HT (Fig. 7). In LT, the situation is reminiscent of lignocellulose degradation by brown rot fungi, which employ Fenton chemistry to selectively metabolize carbohydrates without significant lignin removal, where the extracellular production of Fe²⁺ and H₂O₂ leads to the formation of hydroxyl radicals [84]. It has been proposed that laccases produced in the salivary glands of the host also contribute to delignification during midgut passage in LT [85].

Several lineages of MAGs assigned to *Actinobacteriota* and *Pseudomonadota* encode lytic cellulose monooxygenase (LPMO) and cellobiose dehydrogenase (CDH). LPMOs are highly effective in the depolymerization of recalcitrant polysaccharides because they cleave crystalline cellulose, generating free ends that are accessible to exoglucanases [87, 88]. CDHs shuttle electrons from the oxidation of cellobiose to the catalytic site of LPMO [89]. A metatranscriptomic analysis of a *Labiotermes* sp. has documented that bacterial LPMOs are expressed mainly in the anterior hindgut [11]. It is possible that the oxygen-dependent activities of *Actinomycetales* and *Pseudomonadales* in the hindgut of HT enhance the degradation of wood fibers by polysaccharide-digesting bacteria. However, the small proportion of exoglucanases in the hindgut microbiota of higher termites may have a different explanation. Many endoglucanases, particularly in GH5 and GH9, are processive and have been shown to attack both microcrystalline and amorphous regions of cellulose, producing cellobiose, cellotriose, and cellotetraose as final products [80]. Hence, it is plausible that fiber-associated bacteria can effectively hydrolyze cellulose fibers with the synergistic action of their diverse endoglucanases, as already demonstrated for *Fibrobacter succinogenes*, the most important fiber-degrading bacterium in the rumen [86]. A recent report on a GH5_4 cellulase with high activities on crystalline cellulose from the metagenome of the camel rumen [99] illustrates that further insights into the mechanism of crystalline cellulase degradation by the hindgut bacteria of termites will be gained by integrating both bioinformatics and enzymology.

Both oxidases and oxygenases, and indirectly also peroxidases, require molecular oxygen as co-substrate.
Microsensor measurements have revealed a strong influx of oxygen across the termite gut epithelium, creating microoxic habitats in the periphery of an otherwise anoxic hindgut [90]. Notably, MAGs encoding lignin-modifying enzymes belonged to the same bacterial lineages that were localized at the internal surface of the hindgut wall. Fluorescence in situ hybridization (FISH) has revealed that the microoxic periphery of the hindgut of *Mastotermes darwiniensis* (LT) is colonized by *Alphaproteobacteria* and *Betaproteobacteria* (*Pseudomonadota*) [91]. Clone libraries of the hindgut contents of *Reticulitermes santonensis* and *Reticulitermes speratus* (LT) and a *Nasutitermes* spp. (HT) have identified *Actinomycetota*, *Bacteroidota*, *Pseudomonadota*, and *Spirochaetota* phylotypes in the wall fraction [92–94]. They include the same lineages of *Actinomycetales*, *Corynebacteriales*, and *Propionibacteriales* that are represented by our MAGs with lignin- and cellulose-oxidizing potential (Fig. 7).

## Conclusions

Previous studies revealed unique patterns of bacterial community structure of termites that were attributed to differences in host phylogeny, diet, and microenvironmental factors [95–97]. Our results altogether indicate that this distribution of specific bacterial lineages determines the CAZyme patterns among different host groups. Many members of the bacterial gut microbiota, such as major lineages in the phyla *Elusimicrobiota* and *Patescibacteria*, have a very low content of GHs, indicating they do not contribute to the degradation of polysaccharides and possess only CAZymes necessary for assimilatory functions, such as cell wall maintenance. Many lineages with pronounced (hemi)cellulolytic capacities, such as certain *Bacteroidales* families, occur only in LT, whereas others, such as *Fibrobacterota* and *Actinomycetota*, were present only in HT. In *Spirochaetota*, the fibrolytic capacities differ strongly between lineages from LT and HT, and *Actinomycetota* and *Pseudomonadota* with fibrolytic functions occur only in HT.

Although the reasons for these patterns and their implications for the nutritional symbioses in termite guts seem to be multifactorial and complex, they are ultimately driven by the presence of flagellates in LT and their subsequent loss in HT. In LT, where lignin modification seems to be restricted to host activities in the midgut, the flagellates efficiently depolymerize cellulose but accomplish only incomplete digestion of hemicelluloses, and free-living or flagellate-associated *Bacteroidales* and *Breznakiellaceae* exploit residual cellodextrins and hemicelluloses. In HT, however, the degradation of lignocellulose is partitioned among a consortium of bacterial lineages with different capabilities. *Pseudomonadota* and

*Actinomycetota* that colonize the microoxic periphery of the hindgut partially depolymerize lignin and generate free-ends in the crystalline regions of cellulose using oxygen-dependent activities. This increases the accessibility of the plant fibers for *Fibrobacterota* and HT-specific *Breznakiellaceae*, which possess sophisticated fiber-adhesion mechanisms and effectively depolymerize high-molecular-weight cellulose and hemicelluloses. Finally, the low-molecular remains of (hemi)celluloses released by the activities of the fiber-associated microbiota are processed by various bacterial lineages, including several families of *Clostridia*, which seem to employ hitherto unrecognized strategies involving extracellular cellobiose and cellodextrin phosphorylases.

## Supplementary Information

The online version contains supplementary material available at https://doi.org/10.1186/s40168-024-01917-7.

Additional file 1: Table S1. Termite gut metagenomes metadata. Table S2. MAG classification, metagenomic relative abundance, and quality statistics. Table S3. Functional classification and activity of CAZyme (sub) families represented in the MAGs analyzed in this study. Table S4. Genomic abundance of CAZymes with lignocellulolytic activity in each MAG of the bacterial lineages depicted in Figure 2. Table S5. Genomic abundance of all CAZymes across all MAGs included in this study. Table S6. Prediction of secretion fate for all GH94 genes and the taxonomy of the respective MAGs.

Additional file 2: Figure S1. UMAP biplot of the distribution of CAZymes of termite-associated bacteria color-coded according to phylum in (A) lower termites, (B) higher termites, (C) termite diet, and (D) gut compartment. Figure S2. Distribution of phylogenetic signal across termite-associated bacterial phyla tested against random effects. Figure S3. The drivers of uniqueness in the distribution patterns of (hemi)cellulases and lignin-modification enzymes in 2,223 termite-associated bacterial MAGs. Figure S4. The gene density of cellulases (A) and hemicellulases (B) among the MAGs of *Firmicutes*, *Spirochaetota*, *Fibrobacterota*, and *Bacteroidota* from lower and higher termites. Figure S5. Biplot of the PCA based on the distribution of lignin-modifying enzymes, color coded according to the phylum to which each genome belongs (A) and according to the gut compartment sampled for the metagenomes (B). Figure S6. Distribution of gene densities in CAZymes with pectinase activities in MAGs from different bacterial phyla. The insets break down the two most abundant phyla, *Bacteroidota* and *Planctomycetota*, to the order level.

## Acknowledgements

We thank Sebastian Treitli (Max Planck Institute for Terrestrial Microbiology) for critical discussions and comments on the manuscript.

## Authors' contributions

JFMS and AB conceptualized the study; VH and MAGV performed preliminary analyses; JMFS and VH analyzed the genomic data; JFMS performed statistical analyses; JFMS and AB wrote the manuscript with input from VH and GT. All authors read and approved the final manuscript.

## Funding

Open Access funding enabled and organized by Projekt DEAL. This study was funded by the Max Planck Society and a grant of the Deutsche Forschungs-gemeinschaft (DFG) in the Collaborative Research Center SFB 987. JFMS and MAGV received doctoral fellowships of the International Max Planck Research Schools Principles of Microbial Life (IMPRS-μLife) and Environmental, Cellular, and Molecular Microbiology (IMPRS-Mic) respectively. The funding bodies had
no role in the design of the study and collection, analysis, and interpretation
of data and in writing the manuscript.

**Data availability**

The MAGs used in this study are available in the NCBI Sequence Read
Archive (SRA) under the BioProject accession numbers PRJNA560329 and
PRJNA732531. All data generated or analyzed during this study are included in
this published article and its supplementary information files.

**Declarations**

**Ethics approval and consent to participate**

Not applicable.

**Consent for publication**

Not applicable.

**Competing interests**

The authors declare no competing interests.

Received: 11 June 2024 Accepted: 26 August 2024

Published online: 15 October 2024

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