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1 The origin of fungi-culture in termites was associated with a shift to a mycolytic

2 gut bacteria community

4 Haofu Hu1*, Rafael Rodrigues da Costa1*, Bo Pilgaard2, Morten Schiøtt1, Lene Lange2, and Michael

5 Poulsen1

7 1 Section for Ecology and Evolution, Department of Biology, University of Copenhagen,

8 Universitetsparken 15, 2100 Copenhagen, Denmark.

10 2 Department of Bioengineering, Technical University of Denmark, Søltofts Plads 227, 2800 Kgs.

11 Lyngby, Denmark.

13 *contributed equally to this work.

16 Corresponding author

17 Haofu Hu, Section for Ecology and Evolution, Department of Biology, University of Copenhagen,

18 Universitetsparken 15, 2100 Copenhagen East, Denmark, phone: +4550306440, email:

19 [email protected].

20 Abstract

21 Termites forage on a range of substrates, and it has been suggested that diet shapes the composition

22 and function of termite gut bacterial communities. Through comparative analyses of gut

23 metagenomes in nine termite species with distinct diets, we characterise bacterial community

24 compositions and identify biomass-degrading enzymes and the bacterial taxa that encode them. We

25 find that fungus-growing termite guts are enriched in fungal cell wall-degrading and proteolytic

26 enzymes, while wood-feeding termite gut communities are enriched for plant cell wall-degrading

27 enzymes. Interestingly, wood-feeding termite gut bacteria code for abundant chitinolytic enzymes,

28 suggesting that fungal biomass within the decaying wood likely contributes to gut bacteria or

29 termite host nutrition. Across diets, the dominant biomass-degrading enzymes are predominantly

30 coded for by the most abundant bacterial taxa, suggesting tight links between diet and gut

31 community composition, with the most marked shift being the communities coding for the

32 mycolytic capacity of the fungus-growing termite gut.

34 Key words: Carbohydrate-active enzymes, cellulase, chitinase, HiSeq, metagenome.

35 Introduction

36 Termites are widespread in tropical, subtropical and warm temperate regions (Buczkowski and

37 Bertelsmeir 2016) and form a diverse group of more than 3,000 described species in 281 genera and

38 seven families (Eggleton 2000; 2001; Kambhampati and Eggleton 2000; Inward et al. 2007). They

39 have major impacts on their environments (Buczkowski and Bertelsmeir, 2016), and this success

40 has been attributed to their capacity to use nutritionally-imbalanced, recalcitrant food sources,

41 allowing for colonization of otherwise inaccessible niches (Brune 2014). Different termites forage

42 on distinct substrates, including soil, wood, dung, and fungus (Thongaram et al. 2005; Liu et al.

43 2013), decomposed through intricate interactions with complex gut microbial communities (Brune

44 2014; Brune and Dietrich 2015). In most termites, the main role of gut microbiota is believed to be

45 the digestion of lignocellulose (Ni and Tokuda 2013; Watanabe and Tokuda 2010), but gut

46 microbes also play key roles in nitrogen fixation (Breznak, 1982; Higashi et al. 1992; Sapountzis et

47 al. 2016), microbial defence (e.g., Um et al. 2013) and immune regulation (Round and Mazmanian

48 2009; Viaud et al. 2014), of major importance for the evolutionary history of the symbioses.

50 Approximately 30 million years ago (MYA), the basal higher termite subfamily Macrotermitinae

51 engaged in a mutualistic association with Termitomyces fungi (Aanen et al. 2002; Bourguignon et

52 al. 2015) and shifted composition of the gut microbiota (Dietrich et al. 2014; Otani et al. 2014;

53 2016). Termitomyces decomposes plant material within external fungus gardens (combs) (Nobre et

54 al. 2011; Poulsen et al. 2014), but the gut still remains central in the association, because plant

55 substrate is macerated and mixed with asexual Termitomyces spores in a first gut passage prior to

56 comb deposition (Leuthold et al. 1989). After Termitomyces breaks down the plant substrate, the

57 termites ingest mature parts of the comb in a second gut passage (Leuthold et al. 1989), where gut

58 microbes may contribute enzymes for final digestion of any remaining plant components (Poulsen

59 et al. 2014). This division of labour is consistent with gut bacteria being of importance mainly when

60 the comb material passes through the termite gut in a second passage (cf. Nobre et al. 2011; Poulsen

61 et al. 2014; Poulsen, 2015), but recent work has suggested that partial lignin breakdown may also be

62 accomplished during this first gut passage in Odontotermes formosanus (Li et al. 2017).

64 A set of microbes distinct from the gut microbiota of other termites persists in the fungus-growing

65 termite guts, but limited work has examined the functional implications of this shift (Liu et al. 2013;

66 Poulsen et al. 2014). It has been hypothesized that it was associated with the more protein-rich

67 fungal diet (Dietrich et al. 2014) and/or to break down chitin and other fungal cell wall components

68 (Hyodo et al. 2003; Liu et al. 2013; Poulsen et al. 2014). Termitomyces domestication exposed

69 fungus-growing termite gut communities to large quantities of fungal cell wall glucans (composed

70 of D-glucose monomers), chitin (glucosamine polymer), and glycoproteins (e.g., Reid and

71 Bartnicki-Garcia 1976). Their breakdown requires a combination of carbohydrate-active enzymes

72 (CAZymes; www.cazy.org; Cantarel et al. 2009; Lombard et al. 2014) and fungus-growing termite

73 gut bacteria indeed encode glycoside hydrolase (GH) families of enzymes that may cleave chitin

74 (GH18, GH19, and GH20), beta-glucan (GH55, GH81, and GH128), and alpha-mannan (GH38,

75 GH76, GH92, GH99, and GH125) (Liu et al. 2013; Poulsen et al. 2014).

77 In nature, bacteria are the major chitin degraders and its hydrolysis has been correlated with

78 bacterial abundances in e.g., soil communities (Kielak et al. 2013). In fungus-growing termites,

79 Bacteroidetes and Firmicutes bacteria appear to be the main producers of CAZymes putatively

80 producing mycolytic enzymes (Liu et al. 2013; Poulsen et al. 2014). These studies remained

81 preliminary, however, because they were based on either an unassembled low-coverage

82 metagenome (Liu et al. 2013) or had limited functional predictions (Poulsen et al. 2014). Here, we

83 sequenced the gut metagenome of the fungus-growing termite Odontotermes sp. to elucidate its

84 fungal and plant cell wall-degrading capacities at deeper functional levels (i.e., to EC numbers

85 when possible), assigned putative enzyme functions to gut community members, and performed

86 proteolytic enzyme analyses. To investigate the link between termite diet and gut community

87 composition, we compared our findings to metagenomes from the fungus-growing termite

88 Macrotermes natalensis (Poulsen et al. 2014) and seven non-fungus growing termite species

89 feeding on plant material at different degrees of decomposition; the dung feeder: Amitermes

90 wheeleri (He et al. 2013); two wood feeders: Nasutitermes corniger and Microcerotermes parvus; a

91 litter feeder Cornitermes sp.; two humus feeders: Termes hospes and Neocapritermes taracua; and

92 the soil feeder Cubitermes ugandensis (Rossmassler et al. 2015). We reveal that the shift in gut

93 community composition after the origin of fungus farming was associated with a mycolytic

94 microbiota, providing novel insights into digestion and the role of gut communities in the fungus-

95 growing termite symbiosis.

97 Results

98 Taxonomic composition of fungus-growing termite gut microbiotas

99 We assigned bacterial taxonomies to metagenome contigs by searching for the closest matches of

100 protein-coding genes on each contig against the NR database in NCBI and compared the relative

101 abundance of the contigs in each group to assess the composition of termite gut microbiotas. Ma.

102 natalensis and Odontotermes sp. were distinct from the seven other higher termite species with

103 different diets (Figure 1A), corroborating previous work (Dietrich et al. 2014; Mikaelyan et al.

104 2015) that termites in the same feeding group are similar in gut microbiota composition (Figure 1B,

105 Table S1). Firmicutes and Bacteroidetes dominated both Odontotermes sp. (24% and 25%,

106 respectively) and Ma. natalensis (24% and 30%, respectively) (Figure 1A; Table S1), consistent

107 with 16S rRNA amplicon surveys of these species (Otani et al. 2016) and metagenome analysis of

108 Odontotermes yunnanensis from Southwest China (Liu et al. 2013). Alistipes and Bacteroides were

109 the most abundant genera, representing respectively 5-8% and 2-4% total abundance, in sharp

110 contrast to abundances in the other higher termites (on average 0.1% and 0.5%, respectively) (Table

111 S1). Treponema (Spirochaetes) abundance was generally low in fungus growers and termites that

112 feed on decaying plant material (3 and 6%, respectively) except for the litter feeder Cornitermes sp.

113 (32%), while they were the most abundant taxon in wood feeders (42-45%) (Table S1).

114

115 Fungus- and plant cell wall-targeting enzymes

116 To compare the functional capacity for fungal and plant cell wall degradation, we identified genes

117 belonging to GH families in the CAZy database (Cantarel et al. 2009; Lombard et al. 2014), and

118 classified the genes by their substrate target and thus putative enzyme function by assigning EC

119 numbers (Table 2, S3). Principle Component Analysis (PCA) of GH compositions (Figure 2B)

120 support that gut microbial enzyme capacities are similar between termites with similar diets. This

121 link was also apparent when evaluating relative abundances of GH families coded for by the gut

122 bacteria (Figure 2C). Fungus-growing termite guts were thus systematically different in GH family

123 composition to the guts of termites feeding on plant biomass (Figure 2B, C), reflecting the radical

124 differences in cell wall composition of plant and fungus material.

125

126 GH enzymes targeting fungal cell wall components were enriched in fungus-growing termites, with

127 the most marked differences being for GH125 (34-fold) and GH92 (14-fold) that likely target α-1,6-

128 and 1,2-mannoside (Figure 2C; Table S2). Several β-1,3-glucan-targeting families (GH17, GH128,

129 GH55, and GH87), the chitin-targeting families GH18 and GH19, and lysozyme families with

130 chitin-degradation activities (GH23, GH73, GH24, and GH25), showed 2- to 12-fold enrichment in

131 fungus-growing termites (Figure 2C; Table S2). Similarly, enzymes target galactans that likely

132 belong to fungal cell wall were enriched: GH families targeting β-galactosidase (GH59, GH35, and

133 GH2), along with α-galactosidase and α-N-acetylgalactosaminidase (GH110, GH27, and GH97),

134 showed 3- to 36-fold enrichment compared to non-fungus growing termites. In contrast, and as

135 expected, many GH families encoding genes targeting plant cell wall components were low in

136 relative abundance in fungus-growing termites but higher in wood feeders (Figure 2C; Table S2).

137 Cellulose- (GH94, GH10, GH5) and hemicellulose-targeting families (GH74, GH120, GH39,

138 GH26, GH10, GH11 etc.) were 3- to 5-fold more abundant in wood feeders than in other termites.

139 Interestingly, laccases (EC 1.10.3.2) were only detected in wood feeding termites (Table S3). Lytic

140 polysaccharide monooxygenase (LMPO) were only detected in Cu. ugandensis, where they,

141 however, were present only in very low abundance (Table S3).

142

143 The bacteria encoding the most abundant glycoside hydrolase families

144 To gain further insight into the functional contribution of gut microbiota members to fungal

145 digestion, we grouped enzymes in GH families by fungal cell wall components targeted and

146 bacteria of origin (Table 2; Figure 3). The bacterial orders Clostridiales and Bacteroidales

147 contributed most fungal cell wall-degrading enzymes in fungus-growing termite guts (78%), while

148 Spirochaetales contributed most of these enzymes in the wood-feeding termites (62%; Figure 3).

149 Lysozymes targeting chitin was only abundant in fungus-growing termite gut metagenomes (Table

150 2; Figure 3) and mainly, coded for by members of the Clostridiales. α-mannanase, β-glucanase, and

151 galactosidase targeting mannan/galactomannan and β-glucan in the fungal cell wall, respectively,

152 were also abundant in fungus-growing termite gut, with the majority of these enzymes (60%) being

153 coded for by members of the Bacteroidales (Figure 3), while chitinases and β-N-

154 acetylhexosaminidases were produced by both Clostridiales and Bacteroidales and, to a lesser

155 extent, Spirochaetales in wood feeders. In the remaining termite species, most of the fungal cell

156 wall-degrading enzymes were contributed by Clostridiales, Bacteroidales, and Spirochaetales (17%-

157 92%), but the overall abundance of these enzymes was far lower than in fungus-growing termites

158 and wood feeders (Table 2; Figure 3).

159

160 Distinct protease profiles in termite gut metagenomes

161 We identified proteases in termite gut metagenomes and classified them by catalytic types to

162 investigate the proteolytic potential of gut communities. Proteases were most abundant in fungus-

163 growing termites, which exhibited a 1.4-4 fold enrichment in all catalytic types (Figure 4A),

164 followed by Mi. parvus, Na. corniger (wood-feeders) and A. wheeleri (dung-feeder) (Figure 4B;

165 Table S4). One of the most distinct differences we observed was for glutamic peptidases and mixed

166 peptidases, which were enriched only in the guts of fungus-growing termites. The wood feeder Cu.

167 ugandensis displayed the lowest protease abundance, followed by the litter-feeder Cornitermes sp.

168 As observed for fungal and plant cell wall-degrading enzymes, the most abundant taxa were also

169 predicted to encode for most of the proteases (Figure 4A). Clostridiales and Bacteroidales,

170 Clostridiales and Spirochaetales, and Spirochaetales were the main contributors encoding proteases

171 within the metagenomes of fungus-feeding, dung-feeding, and wood-feeding termites, respectively.

172

173 Discussion

174 Diet is a major driver of taxonomic and functional composition of gut microbial communities

175 (David et al. 2014; Perez-Cobas et al. 2015; Bodawatta et al. 2018). Our characterisation of cell

176 wall degrading enzyme and protease enzymes from higher termites confirm that the functional

177 profiles of gut microbial communities are tight linked with termite diet, consistent with previous

178 work that has focused on community compositions (Otani et al. 2014, 2016; Mikaelyan et al. 2015),

179 and preliminary functional implications of the shift in microbiota structure after the origin of fungus

180 farming in the termite sub-family (Liu et al. 2013; Poulsen et al. 2014). The gut microbiota thus not

181 only functionally complements final plant biomass decomposition by providing oligosaccharide-

182 targeting enzymes (Liu et al. 2013; Poulsen et al. 2014), but also provides key enzymes for the

183 digestion of fungal biomass. The mycolytic potential of the gut microbiota of the South African

184 Odontotermes sp. and Ma. natalensis (this study) was comparable O. yunnanensis from Southwest

185 China (Liu et al. 2013) on, suggesting conserved functions across space and time, and highlighting

186 the robust link between taxonomic and functional ties of the intimate interactions between gut

187 community members and termite hosts.

188

189 Fungus-growing termite gut mycolytic enzymes were primarily coded for by Clostridiales and

190 Bacteroidales, which dominate the core gut microbiota of the Macrotermitinae (Hongoh et al. 2005,

191 Hongoh 2010; Dietrich et al. 2014; Otani et al. 2014; Mikaelyan et al. 2015). The ancestor of the

192 Macrotermitinae likely had a lower termite bacterial gut microbiota (but without protists) (Brune

193 2014; Brune and Dietrich 2015). Fungiculture exposed the gut microbiota to higher amounts of

194 fungal biomass including proteins than to the ancestral lignocellulolytic diet, resulting in a gut

195 microbiota that converged to become more similar to those observed in extant cockroaches

196 (Dietrich, Kohler, and Brune 2014; Otani et al. 2014). This is likely a product of several factors.

197 First, bacterial strains present in the Macrotermitinae ancestor that could utilise a fungal diet were

198 likely selected for and consequently increased in relative abundance (e.g., Alistipes, Dysgonomonas,

199 and members of the Ruminococcaceae) (Dietrich et al. 2014; Otani et al. 2014). Many mycolytic

200 microbes were conceivably already present in termite guts prior to the evolution of fungiculture,

201 since termites feed on partially degraded plant substrates containing fungal biomass. Second,

202 bacteria contributing to the breakdown of lignocellulose in the Macrotermitinae ancestor were likely

203 outcompeted/selected against (e.g., the genus Treponema). Third, novel lineages adopted from other

204 termites or the environment were likely co-opted when fungiculture evolved. This is consistent with

205 recent work demonstrating rampant horizontal transmission of gut bacterial lineages associated with

206 termites (Bourgoignon et al. 2018). Collectively, this led to a gut microbiota enriched in mycolytic

207 (e.g., α-mannanases, β-glucanases, chitinases and proteases) and reduced in lignocellulolytic (e.g.,

208 cellulase, cellobiohydrolase, hemicellulose and laccase) enzymes.

209

210 In sharp contrast to the fungus growers, lignocellulose-degrading enzymes dominated the gut

211 microbiota of wood-feeding termites (Na. corniger and Mi. parvus), as expected from the

212 requirement for the breakdown of recalcitrant plant components, which in fungus farmers are

213 handled by Termitomyces (Poulsen et al. 2014; although lignin cleavage may be initiated during the

214 first gut passage: Li et al. 2017). Interestingly, however, the relatively high abundance of

215 chitinolytic enzymes of Spirochaetales origin (mostly species of the genus Treponema) in wood-

216 feeding termites suggests that the decaying wood these species feed on, harbour fungal biomass that

217 gut bacteria or the termite host may utilise. However, it is also conceivable that fungal cell wall-

218 degrading enzymes, such as β-1,3-glucanase, serve to cleave fungal cell wall to protect against

219 fungal infections (cf., Rosengaus et al. 2014). Experimental work targeting their expression in the

220 presence of fungal pathogens, ideally combined with varying fungal biomass diet content, have the

221 potential to shed light on their relative defensive and dietary roles.

222

223 Proteases were most abundant in fungus-growing termites and mostly coded for by the bacterial

224 taxa contributing mycolytic enzymes. The presence and high abundance of nearly all types of

225 proteases reflects a general and broad proteolytic capacity of the fungus-growing termite gut

226 microbiota that is inevitably related to the protein-rich fungal diet. The presence of proteases in

227 dung and wood feeders could be explained by the presence of fungal biomass in the substrate

228 ingested by these termites. This is consistent with the low abundance of these enzymes in termites

229 feeding on soil and humus with much lower protein content. The only under-represented proteases

230 in fungus-growing termites was threonine peptidases, which has comparable activities as serine and

231 cysteine peptidases (Powers et al. 2002). The absence of these enzymes could thus be compensated

232 for by the presence of other protease families; however, more work will be needed to test what these

233 enzymes indeed target.

234

235 Similarities in host diet have been shown to drive convergence in the functional potential of gut

236 microbes in other organisms (Muegge et al. 2011; Delsuc et al. 2014). Selection for particular

237 physiological traits may, however, not necessarily be directly linked to specific phylogenetic groups

238 of microbes. Exploring communities associated with diverse fungus-growing hosts (and their

239 associated fungi) would allow us to explicitly test for convergent evolution of chitinolytic microbial

240 communities, even if these were likely comprised by different microbial consortia. A number of

241 other insects utilise fungus material as a nutrient source, including fungus-growing ants (Schultz

242 and Brady 2008; Schiøtt et al. 2010), some Drosophila species (Jaenike and James 1991), the

243 Malaysian mushroom-harvesting ant Euprenolepis (Witte and Maschwitz, 2008), Sirex wood wasps

244 (Madden and Coutts, 1979; Hajek et al. 2013), and ambrosia beetles (Batra 1966; Hulcr and

245 Cognato 2010; De Fine Licht and Biedermann 2012). Different bacteria are likely to be involved,

246 but predictions would be that predominantly fungal diets should select for microbial communities

247 with comparable mycolytic capacities. Recent work on mycophagous Drosophila supports that this

248 may be so. Bost et al. (2018) demonstrated that gut bacteria in mycophagous Drosophila are

249 implicated in fungal cell wall metabolism, with cysteine and methionine metabolism enzymes

250 originating from Bacteroidetes, Firmicutes, and Proteobacteria gut microbiota members; phyla that

251 are also abundant and mycolytic in fungus-growing termites. The convergent prevalence of specific

252 protease-producing Firmicutes and Bacteroidetes taxa suggests that they were selected for high-

253 protein host diets, consistent with findings in humans (Eckburg et al. 2005) and pigs (Leser et al.

254 2002), and likely contributing to the observed convergence in fungus-growing termite and

255 cockroach gut metagenomes (Dietrich et al. 2014; Otani et al. 2014; Schauer et al. 2012).

256

257 The prevalence of microbial communities with ample mycolytic capacities in the guts of fungus-

258 growing termite species supports that the shift to a fungal diet was associated with both a functional

259 and compositional shift in gut microbial communities at the onset of fungiculture in termites. The

260 enrichment of GH families encoding fungal cell wall-degrading enzymes and proteases indicates

261 adaptations to the decomposition a fungal diet, consistent with this capacity being absent or less in

262 termites with predominantly plant-based diets. An exception is wood-feeding termites, for which

263 wood-degrading fungi also may comprise an appreciable component of the termite diet. Further

264 work will be needed to elucidate whether the functional capacities of the gut microbiota reflect the

265 amount of fungal biomass in the diet and potential differences in dietary properties of the fungal

266 species fed on. Furthermore, taxonomy assignments were reference-dependent and limited by the

267 closest match present in the databases, with 10-60% of the enzymes identified per metagenome

268 remaining unclassified and most of the classified enzymes remaining unidentified at the genus level,

269 so to improve our understanding of digestive function of gut microbes, further classification of the

270 taxonomic and functional properties of digestive enzymes and nutrient composition of termite diets

271 are needed.

272

273 Material and Methods

274

275 Odontotermes sp. collection

276 Termites from an Odontotermes sp. colony (code: Od127) were collected at the Experimental Farm

277 of the University of Pretoria, South Africa (-25.742700, 28.256517). The species identity of this

278 colony had been previously established as by mitochondrial gene COII barcoding (Otani et al.

279 2014). Fifty old major workers were sampled, and entire guts were dissected and pooled in a 1.5ml

280 Eppendorf tube and stored at –80°C until DNA extraction.

281

282 Gut microbiota DNA extraction

283 Guts were ground in liquid nitrogen, after which DNA was extracted using the Qiagen Animal

284 Tissue Mini-Kit (Qiagen, Hilden, Germany) according to the manufacturer’s description, with the

285 exception that a chloroform extraction step was followed by incubation with protease K. After

286 proteinase K digestion, one volume chloroform/isoamyl alcohol (24/1) was added; tubes were

287 incubated for 15min on a slowly rotating wheel, and centrifuged at 3,000 g for 10 min. The

288 supernatant was transferred to spin columns and the remainder of the manufacturer’s protocol was

289 followed. The quality and purity of samples were determined using NanoDrop® (Thermo

290 Scientific, Wilmington, USA).

291

292 Metagenome sequencing and assembly

293 DNA was sheared to ~350bp fragments, end-repaired, A-tailed, and ligated with Illumina paired-

294 end adaptors (Illumina). The ligated fragments were selected from the desired size on agarose gels

295 and amplified by LM-PCR, and libraries were sequenced with 150bp read lengths on an Illumina

296 HiSeq2500. The quality of raw sequencing reads was assessed before assembly. Reads containing

297 the adaptor, more than 10% N or more than 50% low quality bases (Q-score 5), were removed. To

298 exclude sequences from Termitomyces and the termite hosts, quality-controlled reads were mapped

299 to the Ma. natalensis and Termitomyces genomes (Poulsen et al. 2014) with Burrows-Wheeler

300 Aligner v0.7.15 (Li and Durbin 2010) BWA-MEM algorithm; any aligned reads were filtered.

301

302 Clean reads were assembled by IDBA-UD v1.1.2 (Peng et al. 2010; 2011) with an iterative set up

303 from k-mer size of 19 to 99 at step of 10 (--pre_correction --mink 19 --maxk 99 --step 10).

304 Unassembled reads were picked out by mapping reads back to the initial assembly and assembled

305 separately with the same set up. Redundancies of sequences from the same organism within the

306 metagenome were removed by clustering all contigs at 95% identity with CD-hit v4.6.6 (Li and

307 Godzik 2006) and only the longest contig per cluster was kept. For comparison, high-quality reads

308 of Ma. natalensis old major worker gut microbiota from Poulsen et al. (2014) were also

309 reassembled using the same procedure. Genes in both assemblies were predicted by Prodigal v2.6.3

310 (Hyatt et al. 2010) with metagenomics parameters (-c –p meta).

311

312 Non-fungus growing termite gut metagenomes

313 We obtained seven published non-fungus growing termite gut metagenomes. These were a dung

314 feeder: Amitermes wheeleri (He et al. 2013); two wood feeders: Nasutitermes corniger and

315 Microcerotermes parvus; a litter feeder Cornitermes sp.; two humus feeders: Termes hospes and

316 Neocapritermes taracua; and the soil feeder Cubitermes ugandensis (Rossmassler et al. 2015).

317 Contigs and protein coding genes were downloaded from JGI IMG/M (Table 1).

318

319 Relative abundances of contigs within metagenomes

320 Clean reads of fungus-growing termites gut microbiota were mapped to the contigs by Bowtie

321 v2.2.9 (Langmead and Salzberg 2012), with subsequent masked duplication taking the best match

322 for each read. Contig coverage was estimated by the length and the number of mapped reads per

323 contigs, after which the relative abundance of each contig was calculated as the coverage of the

324 contig divided by the sum of coverage of all contigs (Qin et al. 2010). For non-fungus growing

325 termites, coverage information of contigs was obtained from JGI (Nordberg et al. 2014; Grigoriev et

326 al. 2012) and the relative abundances were estimated in as described for fungus-growing termites.

327

328 Metagenome taxonomic assignment

329 Taxonomic assignment of protein-coding genes was carried out using Diamond (Buchfink et al.

330 2015) alignment against the NR database in NCBI. Alignments with e-values >1e-5 and sequences

331 with identities <30% were removed. Taxonomic information of the top hit was assigned to each

332 gene. Contigs referring to taxonomical levels were determined by a modified lowest common

333 ancestor (LCA)-based algorithm implemented in MEGAN (Huson et al. 2007). Taxonomic

334 classification supported by less than 10% of the genes on each contig were first filtered, and the

335 lowest common ancestor (LCA) for the taxonomic classification of the rest of the genes were

336 assigned to the contig. The relative abundance of contigs belonging to the same taxonomic group

337 was summed up to represent the taxonomic abundance of that taxonomic group in the microbiota.

338 Differences in taxonomic abundances at the phylum level for metagenomes were compared and

339 visualized by Principle Component Analysis (PCA) using R v3.3.2 (R Core Team 2013).

340

341 Carbohydrate-active enzyme analysis

342 We identified genes encoding CAZymes by searching against CAZyme hidden Markov profiles in

343 the dbCAN database (Yin et al. 2012) using HMMer v3.1b (e ≤ 1e-5) (Eddy et al. 1994) and peptide

344 pattern recognition using HotPep (Busk et al. 2013; 2014; 2017). CAZyme family classification

345 supported by both methods was assigned to genes. Genes for which the two methods gave

346 inconsistent results were subjected to an additional Pfam protein domain search using HMMer (e ≤

347 1e-6) to determine the CAZyme family. CAZyme compositional differences between termite

348 species were evaluated using PCA in R. Enzyme functions of genes in each CAZyme family were

349 determined by BLASTp searches (e ≤ 1e-5) against the ExPASy enzyme records, CCD searches

350 against COG database (Marchler-Bauer et al. 2017) and GhostKOALA searches using the KEGG

351 database online tool (Kanehisa et al. 2016). Genes coding for enzymes related to fungal and plant

352 cell wall degradation were selected and their substrate targets and bacterial taxonomy assigned and

353 manually checked.

354

355 Protease identification and analyses

356 Protein sequences of annotated genes in metagenomes were aligned to MEROPS database

357 (Rawlings et al. 2018) by BLASTp (e ≤ 1e-5) to identify proteases. Genes that matched at least 50%

358 of the sequences in the database were selected. The catalytic type and protease family classification

359 of genes were annotated based on their most similar matches in the MEROPS database, and

360 bacterial taxonomy was assigned and manually checked as described for the CAZyme analysis.

361

362 Data deposition

363 Clean reads and metagenome assembly have been submitted to the SRA and GenBank under

364 BioProject accession numbers PRJNA476694 and PRJNA193472.

365

366 Conflict of interest

367 The authors declare no conflict of interest.

368

369 Acknowledgements

370 We thank Z. Wilhelm de Beer, Michael J. Wingfield, and the staff and students at the Forestry and

371 Agricultural Biotechnology Institute, University of Pretoria, for hosting field work and contributed

372 to the research environment, and Christine Beemelmanns, René Benndorf, Saria Otani, Margo

373 Wisselink, Sabine M. E. Vreeburg, Duur K. Aanen, Lennart van de Peppel, Nina Kreuzenbeck, and

374 Victoria L. Challinor for help with excavations. This study was funded by the CAPES Foundation,

375 Ministry of Education of Brazil (grant BEX: 13240/13-7) to RRdC and a Villum Kann Rasmussen

376 Young Investigator Fellowship (VKR10101) to MP.

377

378 Author Contributions

379 HH and RRdC designed the experiments and analyses, collected material in the field, extracted

380 DNA, sent samples for sequencing, and drafted the first version of the figures, tables, and

381 manuscript. HH carried out the assembly of the metagenome and the functional analyses. BP and

382 LL carried out HotPep analyses and contributed to data interpretation. MS contributed to data

383 analyses and interpretations. MP supervised HH and RRdC, helped design the study, contributed

384 with comments on analyses and the first versions of figures, tables, and text. All authors contributed

385 to writing the manuscript.

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699 Tables with legends 700 701 Table 1. Summary of metagenome data.

Sample Assembly data Assembly Number Termite species Diet Reference Accession No. size size N50 (bp) of genes (Mbp) (Gbp) Odontotermes sp. Fungus NA 8.6 PRJNA476694 797.15 1,133 753,265 Macrotermes natalensis Fungus Poulsen et al. 2014 8.5 PRJNA193472 498.71 1,569 486,207 Amitermes wheeleri Dung He et al. 2013 0.3 IMG:2030936000 139.3 512 321,461 Nasutitermes corniger Wood Rossmassler et al. 2015 46.8 IMG:3300002119 634.52 980 1,338,688 Microcerotermes parvus Wood Rossmassler et al. 2015 43.2 IMG:3300002449 712.35 945 1,419,719 Cornitermes sp. Litter Rossmassler et al. 2015 45.9 IMG:3300002450 131.61 519 2,945,692 Termes hospes Humus Rossmassler et al. 2015 34.2 IMG:3300002469 1212.37 424 2,975,319 Neocapritermes taracua Humus Rossmassler et al. 2015 28.3 IMG:3300002505 885.58 454 2,115,406 Cubitermes ugandensis Soil Rossmassler et al. 2015 32.0 IMG:3300002127 1121.61 475 3,061,751 702 703

Table 2. Relative abundance of fungal cell wall degrading enzymes in the gut metagenomes of nine termites with different diets. Mn: Macrotermes natalensis, Od: Odontotermes sp., Nc: Nasutitermes corniger, Aw: Amitermes wheeleri, Mp: Microcerotermes parvus, Co: Cornitermes sp., Th: Termes hospes, Nt: Neocapritermes taracua, Cu: Cubitermes ugandensis. For the full results, see Table S2. Fungus Soil Dung Humus Litter Wood Substrate Enzyme activity EC Mn Od Cu Aw Nt Th Co Mp Nc Chitin Chitinase 3.2.1.14 GH18, GH19 2.34E-04 5.52E-05 4.15E-06 2.73E-05 8.58E-06 1.30E-05 2.43E-05 7.05E-05 7.85E-05 GH24, GH25, Lysozyme 3.2.1.17 2.57E-04 9.23E-05 1.60E-05 9.21E-06 1.86E-05 1.18E-05 8.55E-06 1.05E-05 1.57E-05 GH73, GH23, GH22 β-N-acetylhexosaminidase 3.2.1.52 GH3, GH20, GH18 3.21E-04 2.14E-04 2.53E-05 1.73E-04 9.34E-05 7.65E-05 1.41E-04 3.93E-04 3.68E-04 Chitosan Chitin deacetylase 3.5.1.41 CE4 4.19E-06 7.27E-06 2.09E-06 1.69E-05 8.71E-06 3.55E-06 1.19E-05 2.68E-05 6.72E-06 Chitosanase 3.2.1.132 - 0 0 0 0 0 0 0 0 0 exo-β-glucosaminidase 3.2.1.165 GH2 3.61E-05 1.97E-05 1.97E-06 0 5.74E-06 1.83E-06 0.00E+00 1.51E-06 2.76E-07 Mannan endo-1,2-α-mannanase 3.2.1.- GH99 8.87E-05 9.04E-05 8.78E-06 0 2.96E-05 2.50E-05 1.53E-05 3.46E-05 3.43E-05 exo-α-mannosidase 3.2.1.24 GH38, GH92 2.97E-04 1.60E-04 3.66E-05 4.33E-05 1.22E-04 6.57E-05 1.69E-05 5.19E-05 2.75E-05 endo-1,6-α-mannanase 3.2.1.101 GH76 2.79E-05 1.27E-05 1.41E-06 0 4.32E-06 2.49E-06 0 0 0 exo-1,6-α-mannosidase 3.2.1.- GH125 3.98E-05 3.44E-05 1.13E-06 0 1.07E-06 2.10E-06 1.40E-07 1.14E-06 1.95E-06 β Glucan endo-1,3-β-glucanase 3.2.1.39 GH16 4.72E-05 3.63E-05 1.75E-06 0 6.83E-07 2.35E-06 4.05E-06 2.75E-05 1.31E-05 exo-1,3-β-glucanase 3.2.1.58 GH55, GH17, GH5 2.30E-05 1.81E-05 3.36E-06 0 5.06E-06 3.35E-06 3.24E-06 3.40E-05 5.53E-06 endo-1,6-β-glucosidase 3.2.1.75 GH30 4.86E-05 1.63E-05 6.09E-07 6.14E-06 4.52E-06 3.45E-06 0 1.34E-06 0 exo-β-glucosidase 3.2.1.21 GH13 4.66E-04 3.28E-04 6.78E-06 1.41E-04 5.62E-05 6.42E-05 1.07E-04 3.13E-04 2.67E-04 α glucan endo-1,3-α-glucosidase 3.2.1.59 GH71 0 0 0 0 0 2.78E-07 0 0 0 exo-1,3-α-glucanase 3.2.1.84 GH31 0 0 0 0 0 0 0 0 0 endo-1,4-α-glucanase 3.2.1.20 GH13, GH31, GH4 1.96E-05 4.48E-05 1.02E-05 6.14E-06 1.07E-05 1.03E-05 5.51E-06 4.00E-05 5.75E-05 exo-1,4-α-glucanase 3.2.1.3 GH3, GH1 2.37E-05 2.52E-05 1.25E-06 0 2.44E-06 1.84E-06 3.77E-07 2.28E-06 3.12E-07 Fungal galactan GH27, GH110, exo-α-galactosidase 3.2.1.22 GH97 2.37E-04 1.95E-04 2.21E-05 1.39E-04 8.47E-05 8.44E-05 7.54E-05 1.53E-04 1.71E-04 β-galactosidase 3.2.1.23 GH2, GH42, GH35 5.70E-04 4.49E-04 1.90E-05 3.13E-05 7.89E-05 9.04E-05 1.09E-04 2.11E-04 2.22E-04 α-N-acetylgalactosaminidase 3.2.1.49 GH109, GH27 1.40E-04 8.51E-05 2.30E-05 7.37E-06 1.08E-04 7.06E-05 1.03E-05 1.10E-05 2.58E-06 Sum 2.88E-03 1.88E-03 1.86E-04 6.00E-04 6.43E-04 5.33E-04 5.33E-04 1.38E-03 1.27E-03

Figures

Figure 1.

Figure 2.

Figure 3.

Figure 4.

30 bioRxiv preprint doi: https://doi.org/10.1101/440172; this version posted October 11, 2018. The copyright holder for this preprint (which was not certified by peer review) is the author/funder, who has granted bioRxiv a license to display the preprint in perpetuity. It is made available under aCC-BY 4.0 International license.

Figure legends

Figure 1. (A) Relative abundances of bacterial phyla each comprising more than 1% of the microbiota in the guts of termite species with different diets. Termite species are arranged by the degree of plant degradation in the diet. (B) Principal component analysis of community similarities of termites with different diet. Mn: Macrotermes natalensis, Od: Odontotermes sp., Nc:

Nasutitermes corniger, Aw: Amitermes wheeleri, Mp: Microcerotermes parvus, Co: Cornitermes sp., Th: Termes hospes, Nt: Neocapritermes taracua, Cu: Cubitermes ugandensis.

Figure 2. (A) Simplified schematic of fungal (top) and plant (bottom) cell wall structures and targets of enzymes identified in the metagenomes. (B) Principal component analysis (PCA) of relative abundances of carbohydrate-active enzymes in worker gut metagenomes; shapes represent termite feeding groups. Mn: Macrotermes natalensis, Od: Odontotermes sp., Nc: Nasutitermes corniger, Aw: Amitermes wheeleri, Mp: Microcerotermes parvus, Co: Cornitermes sp., Th: Termes hospes, Nt: Neocapritermes taracua, Cu: Cubitermes ugandensis. (C) Relative abundances of enriched (red) and contracted (blue) GH families across the nine metagenomes shown as Log2 fold changes. The symbols on the left of the heatmap follow the legend from Figure 2A, and indicate which predicted substrates that are targeted by the enzymes by members of a given GH family, and family names of GH families with the highest average abundances across all metagenomes are indicated on the right.

Figure 3. The relative abundance of fungal cell wall degrading enzymes encoded by the top five most abundant bacterial orders in the nine termite species. Mn: Macrotermes natalensis, Od:

Odontotermes sp., Nc: Nasutitermes corniger, Aw: Amitermes wheeleri, Mp: Microcerotermes parvus, Co: Cornitermes sp., Th: Termes hospes, Nt: Neocapritermes taracua, Cu: Cubitermes

31 bioRxiv preprint doi: https://doi.org/10.1101/440172; this version posted October 11, 2018. The copyright holder for this preprint (which was not certified by peer review) is the author/funder, who has granted bioRxiv a license to display the preprint in perpetuity. It is made available under aCC-BY 4.0 International license.

ugandensis.

Figure 4. (A) The relative abundances of proteases encoded for by the five most contributing bacterial orders across the nine termite species. (B) A heatmap of the relative abundances of the most abundant proteases across the nine species shown as Log2 fold change (red: enriched and blue: contracted). Mn: Macrotermes natalensis, Od: Odontotermes sp., Nc: Nasutitermes corniger,

Aw: Amitermes wheeleri, Mp: Microcerotermes parvus, Co: Cornitermes sp., Th: Termes hospes,

Nt: Neocapritermes taracua, Cu: Cubitermes ugandensis.

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Supplementary Material for

The origin of fungiculture in termites was associated with a shift to a mycolytic gut bacteria community

Haofu Hu1*, Rafael Rodrigues da Costa1*, Bo Pilgaard2, Morten Schiøtt1, Lene Lange2, and Michael

Poulsen.1

1Section for Ecology and Evolution, Department of Biology, University of Copenhagen,

Universitetsparken 15, 2100 Copenhagen, Denmark.

2 Department of Chemical and Biochemical Engineering, Technical University of Denmark, Søltofts

Plads, 2800 Kgs. Lyngby, Denmark.

* contributed equally to this work.

Table S1. Bacterial community composition of metagenome contigs of gut metagenomes of nine termites with different diets at the phylum, class, order, family, and genus levels. Taxonomy classifications with more than 0.1% of the total abundance are shown. Mn: Macrotermes natalensis, Od: Odontotermes sp., Nc: Nasutitermes corniger, Aw: Amitermes wheeleri, Mp: Microcerotermes parvus, Co: Cornitermes sp., Th: Termes hospes, Nt: Neocapritermes taracua, Cu: Cubitermes ugandensis.

Level Taxa Mn Od Cu Aw Nt Th Co Mp Nc Spirochaetes 0.0343 0.0688 0.0095 0.2114 0.1089 0.0841 0.3359 0.5026 0.4624 Firmicutes 0.2593 0.2361 0.0912 0.2958 0.2213 0.2512 0.1625 0.1285 0.1219 Bacteroidetes 0.2996 0.2488 0.1259 0.0423 0.1455 0.0888 0.0382 0.0487 0.0412

Proteobacteria 0.0791 0.0763 0.0933 0.0683 0.0955 0.1143 0.0326 0.0398 0.0426

Fibrobacteres 0.0005 0.0059 0.0014 0.0151 0.0008 0.0014 0.0454 0.0795 0.1291 Planctomycetes 0.0325 0.0167 0.0165 0.0102 0.0346 0.0555 0.0143 0.0033 0.0020

Actinobacteria 0.0105 0.0194 0.0074 0.0084 0.0219 0.0082 0.0033 0.0048 0.0033 Phylum Synergistetes 0.0083 0.0059 0.0012 0.0162 0.0071 0.0175 0.0008 0.0007 0.0011 Acidobacteria 0.0010 0.0006 0.0009 0.0040 0.0121 0.0027 0.0005 0.0047 0.0081 Other 0.0211 0.0234 0.0349 0.0485 0.0417 0.0425 0.0235 0.0394 0.0360 Unassigned 0.2539 0.2981 0.6177 0.2799 0.3105 0.3338 0.3431 0.1481 0.1521 Spirochaetia 0.033 0.067 0.008 0.206 0.105 0.080 0.331 0.495 0.454 Clostridia 0.230 0.206 0.072 0.245 0.186 0.212 0.129 0.095 0.092 Bacteroidia 0.261 0.208 0.069 0.023 0.099 0.055 0.019 0.027 0.019 Deltaproteobacteria 0.028 0.030 0.026 0.035 0.063 0.049 0.012 0.015 0.018 Bacilli 0.011 0.013 0.011 0.029 0.017 0.021 0.015 0.015 0.013 Betaproteobacteria 0.017 0.017 0.043 0.009 0.009 0.033 0.004 0.005 0.005 0.000 0.003 0.000 0.006 0.000 0.001 0.006 0.039 0.073

Fibrobacteria Gammaproteobacteria 0.013 0.010 0.015 0.015 0.013 0.017 0.010 0.012 0.011

Chitinispirillia 0.000 0.000 0.000 0.008 0.000 0.000 0.033 0.026 0.029 Class Planctomycetia 0.011 0.008 0.009 0.006 0.019 0.027 0.008 0.002 0.001 Actinobacteria 0.006 0.015 0.004 0.007 0.013 0.006 0.002 0.004 0.002 Synergistia 0.008 0.006 0.001 0.016 0.007 0.018 0.001 0.001 0.001 Planctomycetes_bacterium_RBG_16_64_12 0.003 0.002 0.003 0.002 0.007 0.010 0.003 0.000 0.000 Fibrobacteres_bacterium_CG2_30_45_31 0.000 0.002 0.000 0.001 0.000 0.000 0.001 0.007 0.012 Other 0.060 0.078 0.089 0.110 0.128 0.106 0.058 0.084 0.081 Unassigned 0.318 0.336 0.649 0.283 0.333 0.364 0.369 0.174 0.190 Spirochaetales 0.033 0.066 0.203 0.007 0.104 0.079 0.329 0.492 0.451 Clostridiales 0.223 0.200 0.237 0.068 0.179 0.205 0.124 0.092 0.090 Bacteroidales 0.259 0.205 0.023 0.067 0.096 0.054 0.019 0.026 0.018 Fibrobacterales 0.000 0.003 0.006 0.000 0.000 0.001 0.006 0.040 0.073 Bacillales 0.009 0.010 0.024 0.008 0.014 0.015 0.013 0.013 0.011 Desulfovibrionales 0.014 0.013 0.015 0.005 0.033 0.007 0.003 0.003 0.005

Chitinispirillales 0.000 0.000 0.008 0.000 0.000 0.000 0.033 0.026 0.029 Planctomycetales 0.010 0.007 0.005 0.009 0.018 0.026 0.007 0.001 0.001 Synergistales 0.008 0.006 0.016 0.001 0.007 0.018 0.001 0.001 0.001 Order Myxococcales 0.001 0.001 0.001 0.011 0.001 0.028 0.003 0.001 0.001 Rhodocyclales 0.001 0.003 0.001 0.012 0.003 0.020 0.001 0.001 0.000 Candidatus_Adiutrix_intracellularis 0.003 0.006 0.007 0.000 0.012 0.002 0.002 0.003 0.003 Planctomycetes_bacterium_RBG_16_64_12 0.003 0.002 0.002 0.003 0.007 0.011 0.003 0.000 0.000 Fibrobacteres_bacterium_CG2_30_45_31 0.000 0.002 0.001 0.000 0.000 0.000 0.001 0.006 0.012 Other 0.099 0.121 0.167 0.134 0.180 0.159 0.083 0.117 0.111 Unassigned 0.336 0.356 0.284 0.673 0.344 0.375 0.373 0.179 0.194

Spirochaetaceae 0.032 0.066 0.007 0.204 0.104 0.079 0.329 0.492 0.452

Ruminococcaceae 0.036 0.049 0.013 0.074 0.040 0.056 0.019 0.017 0.017 Clostridiaceae 0.026 0.029 0.016 0.050 0.034 0.030 0.020 0.020 0.019

Family Lachnospiraceae 0.025 0.021 0.013 0.045 0.022 0.027 0.019 0.018 0.018 Porphyromonadaceae 0.040 0.049 0.022 0.005 0.044 0.019 0.005 0.008 0.004

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Rikenellaceae 0.107 0.065 0.002 0.001 0.003 0.002 0.001 0.001 0.001 Fibrobacteraceae 0.000 0.003 0.000 0.006 0.000 0.001 0.006 0.039 0.073 Bacteroidaceae 0.043 0.024 0.009 0.005 0.013 0.009 0.004 0.004 0.003 Chitinispirillaceae 0.000 0.000 0.000 0.008 0.000 0.000 0.033 0.026 0.029 Desulfovibrionaceae 0.013 0.012 0.004 0.014 0.032 0.006 0.002 0.003 0.005 Peptococcaceae 0.008 0.011 0.008 0.016 0.017 0.013 0.006 0.007 0.006 Planctomycetaceae 0.007 0.005 0.006 0.004 0.014 0.022 0.006 0.001 0.000 Paenibacillaceae 0.004 0.005 0.004 0.012 0.006 0.007 0.006 0.007 0.005 Synergistaceae 0.008 0.006 0.001 0.015 0.007 0.017 0.001 0.001 0.001 Eubacteriaceae 0.003 0.003 0.003 0.015 0.008 0.006 0.004 0.004 0.003 Rhodocyclaceae 0.001 0.003 0.012 0.001 0.003 0.021 0.001 0.001 0.000 Candidatus_Adiutrix_intracellularis 0.003 0.006 0.000 0.007 0.012 0.002 0.002 0.004 0.003 Planctomycetes_bacterium_RBG_16_64_12 0.003 0.002 0.003 0.002 0.007 0.010 0.003 0.000 0.000 Fibrobacteres_bacterium_CG2_30_45_31 0.000 0.002 0.000 0.001 0.000 0.000 0.001 0.007 0.012 Archangiaceae 0.000 0.000 0.006 0.000 0.000 0.014 0.001 0.000 0.000 Other 0.143 0.164 0.171 0.225 0.244 0.211 0.108 0.145 0.135 Unassigned 0.496 0.476 0.699 0.290 0.388 0.448 0.423 0.196 0.214 Other 0.216 0.249 0.232 0.358 0.345 0.321 0.159 0.195 0.181 Treponema 0.031 0.064 0.007 0.197 0.102 0.077 0.322 0.482 0.444 Alistipes 0.083 0.049 0.002 0.001 0.002 0.002 0.001 0.001 0.000 Bacteroides 0.042 0.024 0.009 0.005 0.013 0.009 0.003 0.004 0.003 Clostridium 0.019 0.022 0.012 0.039 0.026 0.022 0.016 0.016 0.015 Dysgonomonas 0.018 0.010 0.003 0.001 0.005 0.003 0.001 0.001 0.001 Desulfovibrio 0.010 0.009 0.004 0.012 0.028 0.005 0.002 0.003 0.004 Parabacteroides 0.006 0.009 0.005 0.001 0.013 0.006 0.001 0.002 0.000 Candidatus Adiutrix 0.003 0.006 0.000 0.008 0.012 0.002 0.002 0.004 0.003 Ruminococcus 0.007 0.004 0.004 0.027 0.008 0.009 0.003 0.005 0.005

Genus Ruminiclostridium 0.003 0.004 0.003 0.019 0.006 0.007 0.004 0.006 0.005 Fibrobacter 0.000 0.003 0.000 0.006 0.000 0.001 0.006 0.040 0.073 Eubacterium 0.002 0.002 0.002 0.013 0.006 0.005 0.003 0.003 0.003 Planctomycetes_bacterium_RBG_16_64_12 0.003 0.002 0.003 0.002 0.007 0.010 0.003 0.000 0.000 Fibrobacteres_bacterium_CG2_30_45_31 0.000 0.002 0.000 0.001 0.000 0.000 0.001 0.007 0.012 Sporobacter 0.000 0.001 0.001 0.011 0.014 0.012 0.005 0.001 0.001 Chitinispirillum 4.5E-05 0.000143 0.00048 0.00797 0.00016 0.00021 0.033 0.02568 0.02858 Azovibrio 5.26E-05 3.77E-05 0.00081 0.00021 0.00012 0.01417 4E-05 0.00024 1.1E-05 Unassigned 0.555 0.540 0.712 0.293 0.411 0.496 0.434 0.205 0.221

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Table S2. Relative abundance of glycoside hydrolase families in gut metagenomes of nine species of termites with different diets. Mn: Macrotermes natalensis, Od: Odontotermes sp., Nc: Nasutitermes corniger, Aw: Amitermes wheeleri, Mp: Microcerotermes parvus, Co: Cornitermes sp., Th: Termes hospes, Nt: Neocapritermes taracua, Cu: Cubitermes ugandensis.

Fold change between Glycoside fungus vs. Target cell wall Hydrolase Mn Od Mp Nc Cu Th Nt Co Aw Enzyme List non-fungus component family growing termites β-galactosidase (EC 3.2.1.23); GH59 7.09E-06 1.30E-05 0.00E+00 0.00E+00 1.70E-07 3.99E-07 1.37E-06 0.00E+00 0.00E+00 36.23 galactose galactocerebrosidase (EC 3.2.1.46) GH125 3.98E-05 3.45E-05 1.14E-06 1.95E-06 1.13E-06 2.10E-06 1.07E-06 1.40E-07 0.00E+00 34.57 mannan exo-α-1,6-mannosidase (EC 3.2.1.-) GH108 7.23E-05 8.95E-05 1.99E-06 2.65E-06 7.68E-07 2.49E-06 3.79E-06 1.23E-06 9.21E-06 25.58 chitin lysozyme (EC 3.2.1.17) glucoamylase (EC 3.2.1.3); α-glucosidase (EC GH97 1.45E-04 1.03E-04 4.56E-06 2.64E-06 4.77E-06 6.68E-06 1.58E-05 2.52E-06 0.00E+00 23.41 α-glucan 3.2.1.20); α-galactosidase (EC 3.2.1.22) GH89 4.22E-05 1.52E-05 4.14E-07 3.75E-06 1.33E-06 2.23E-07 4.37E-06 9.42E-08 0.00E+00 19.73 NA α-N-acetylglucosaminidase (EC 3.2.1.50) endo-α-1,4-polygalactosaminidase (EC GH114 6.17E-06 1.19E-05 5.49E-07 1.72E-07 0.00E+00 1.15E-07 2.08E-06 6.79E-07 0.00E+00 17.59 NA 3.2.1.109) mannosyl-oligosaccharide α-1,2-mannosidase (EC 3.2.1.113); mannosyl-oligosaccharide α- 1,3-mannosidase (EC 3.2.1.-); mannosyl- oligosaccharide α-1,6-mannosidase (EC GH92 2.88E-04 1.52E-04 1.13E-05 7.80E-06 1.01E-05 1.77E-05 5.54E-05 3.25E-06 0.00E+00 14.58 mannan 3.2.1.-); α-mannosidase (EC 3.2.1.24); α-1,2- mannosidase (EC 3.2.1.-); α-1,3-mannosidase (EC 3.2.1.-); α-1,4-mannosidase (EC 3.2.1.-); mannosyl-1-phosphodiester α-1,P- mannosidase (EC 3.2.1.-) GH25 2.23E-04 8.70E-05 1.70E-05 1.48E-05 9.91E-06 8.83E-06 2.33E-05 9.36E-06 6.14E-06 12.14 chitin lysozyme (EC 3.2.1.17) mycodextranase (EC 3.2.1.61); α-1,3- GH87 1.56E-05 6.66E-06 2.97E-06 9.58E-07 0.00E+00 7.36E-07 2.11E-06 1.25E-06 0.00E+00 9.72 α-glucan glucanase (EC 3.2.1.59) GH24 6.17E-05 6.08E-05 2.39E-06 4.94E-06 8.34E-06 1.12E-05 5.12E-06 3.19E-06 1.54E-05 8.49 chitin lysozyme (EC 3.2.1.17) exo-β-1,3-glucanase (EC 3.2.1.58); endo-β- GH55 5.90E-05 2.66E-05 2.11E-05 0.00E+00 7.01E-07 4.09E-06 8.95E-06 3.09E-06 0.00E+00 7.89 β-glucan 1,3-glucanase (EC 3.2.1.39) GH133 1.16E-04 1.05E-04 5.24E-06 3.79E-06 1.21E-05 1.59E-05 2.51E-05 3.69E-06 5.37E-05 6.49 NA amylo-α-1,6-glucosidase (EC 3.2.1.33); α-L-fucosidase (EC 3.2.1.51); α-1,2-L- GH95 1.60E-04 1.35E-04 4.65E-05 4.21E-05 6.80E-06 2.44E-05 3.20E-05 1.37E-05 0.00E+00 6.22 NA fucosidase (EC 3.2.1.63); α-L-galactosidase (EC 3.2.1.-) GH93 1.44E-05 1.50E-05 3.54E-06 3.89E-06 4.50E-07 2.55E-06 4.10E-06 2.59E-06 0.00E+00 6.01 NA exo-α-L-1,5-arabinanase (EC 3.2.1.-) GH110 1.54E-05 9.71E-06 6.69E-07 4.68E-07 0.00E+00 3.22E-06 9.24E-06 1.08E-06 0.00E+00 5.98 galactose α-galactosidase (EC 3.2.1.22); α-1,3- 36 bioRxiv preprint doi: https://doi.org/10.1101/440172; this version posted October 11, 2018. The copyright holder for this preprint (which was not certified by peer review) is the author/funder, who has granted bioRxiv a license to display the preprint in perpetuity. It is made available under aCC-BY 4.0 International license.

galactosidase (EC 3.2.1.-) invertase (EC 3.2.1.26); endo-inulinase (EC 3.2.1.7); β-2,6-fructan 6-levanbiohydrolase (EC 3.2.1.64); endo-levanase (EC 3.2.1.65); exo-inulinase (EC 3.2.1.80); fructan β-(2,1)- fructosidase/1-exohydrolase (EC 3.2.1.153); fructan β-(2,6)-fructosidase/6-exohydrolase (EC 3.2.1.154); sucrose:sucrose 1- fructosyltransferase (EC 2.4.1.99); GH32 5.32E-05 6.39E-05 5.21E-06 2.90E-06 5.14E-06 9.92E-06 1.56E-05 5.09E-06 2.64E-05 5.83 NA fructan:fructan 1-fructosyltransferase (EC 2.4.1.100); sucrose:fructan 6- fructosyltransferase (EC 2.4.1.10); fructan:fructan 6G-fructosyltransferase (EC 2.4.1.243); levan fructosyltransferase (EC 2.4.1.-); [retaining] sucrose:sucrose 6- fructosyltransferase (6-SST) (EC 2.4.1.-); cycloinulo-oligosaccharide fructanotransferase (EC 2.4.1.-) GH128 7.74E-06 4.01E-05 8.46E-06 1.01E-05 5.63E-07 3.09E-06 1.02E-06 6.18E-06 0.00E+00 5.70 β-glucan β-1,3-glucanase (EC 3.2.1.39) processing α-glucosidase (EC 3.2.1.106); α- 1,3-glucosidase (EC 3.2.1.84); α-glucosidase GH63 3.42E-05 8.44E-06 1.56E-05 3.61E-06 0.00E+00 1.32E-06 3.42E-06 2.51E-06 0.00E+00 5.64 α-glucan (EC 3.2.1.20); mannosylglycerate α- mannosidase / mannosylglycerate hydrolase (EC 3.2.1.170) peptidoglycan lytic transglycosylase (EC GH104 7.94E-07 2.13E-06 1.87E-07 0.00E+00 1.08E-06 3.77E-07 1.44E-07 6.00E-08 0.00E+00 5.54 NA 3.2.1.-) d-4,5-unsaturated β-glucuronyl hydrolase (EC GH88 1.25E-04 7.38E-05 2.45E-05 1.34E-05 1.67E-06 1.45E-05 1.35E-05 1.17E-05 4.73E-05 5.51 NA 3.2.1.-) β-galactosidase (EC 3.2.1.23); exo-β- glucosaminidase (EC 3.2.1.165); exo-β-1,4- GH35 3.90E-05 2.84E-05 1.10E-05 1.38E-05 1.60E-06 7.81E-06 5.55E-06 7.23E-06 0.00E+00 5.02 galactose galactanase (EC 3.2.1.-); β-1,3-galactosidase (EC 3.2.1.-) β-galactosidase (EC 3.2.1.23) ; β-mannosidase (EC 3.2.1.25); β-glucuronidase (EC 3.2.1.31); α-L-arabinofuranosidase (EC 3.2.1.55); GH2 6.24E-04 4.34E-04 2.23E-04 1.77E-04 1.82E-05 1.06E-04 9.71E-05 1.04E-04 2.12E-05 4.96 galactose/mannan mannosylglycoprotein endo-β-mannosidase (EC 3.2.1.152); exo-β-glucosaminidase (EC 3.2.1.165) α-L-rhamnosidase (EC 3.2.1.40); GH78 1.95E-04 1.84E-04 8.08E-05 2.42E-05 2.26E-05 4.37E-05 8.90E-05 1.32E-05 2.46E-05 4.45 NA rhamnogalacturonan α-L-rhamnohydrolase (EC 3.2.1.174) glycoprotein endo-α-1,2-mannosidase (EC GH99 9.14E-05 9.21E-05 3.46E-05 3.43E-05 8.77E-06 2.50E-05 2.96E-05 1.52E-05 0.00E+00 4.36 mannan 3.2.1.130); mannan endo-1,2-α-mannanase (3.2.1.-)

37 bioRxiv preprint doi: https://doi.org/10.1101/440172; this version posted October 11, 2018. The copyright holder for this preprint (which was not certified by peer review) is the author/funder, who has granted bioRxiv a license to display the preprint in perpetuity. It is made available under aCC-BY 4.0 International license.

β-N-acetylgalactosaminidase (EC 3.2.1.53); GH123 2.78E-05 1.75E-05 7.69E-06 1.61E-06 1.60E-06 1.54E-05 9.78E-06 3.05E-06 0.00E+00 4.06 NA glycosphingolipid β-N- acetylgalactosaminidase (EC 3.2.1.-) unsaturated rhamnogalacturonyl hydrolase GH105 1.83E-04 1.63E-04 8.54E-05 5.54E-05 2.54E-06 4.46E-05 2.92E-05 3.68E-05 4.88E-05 4.00 NA (EC 3.2.1.172); d-4,5-unsaturated β- glucuronyl hydrolase (EC 3.2.1.-) glucan endo-1,3-β-glucosidase (EC 3.2.1.39); glucan 1,3-β-glucosidase (EC 3.2.1.58); GH17 4.29E-06 4.18E-06 0.00E+00 1.53E-07 1.83E-06 1.56E-06 3.68E-06 3.56E-07 0.00E+00 3.91 β-glucan licheninase (EC 3.2.1.73); ABA-specific β- glucosidase (EC 3.2.1.175); β-1,3- glucanosyltransglycosylase (EC 2.4.1.-) lacto-N-biose phosphorylase or galacto-N- biose phosphorylase (EC 2.4.1.211); D- GH112 3.23E-05 2.13E-05 1.28E-05 1.14E-05 0.00E+00 1.24E-05 9.93E-07 1.09E-05 0.00E+00 3.87 NA galactosyl-β-1,4-L-rhamnose phosphorylase (EC 2.4.1.247); galacto-N-biose/lacto-N-biose phosphorylase (EC 2.4.1.-) polygalacturonase (EC 3.2.1.15); α-L- rhamnosidase (EC 3.2.1.40); exo- polygalacturonase (EC 3.2.1.67); exo- polygalacturonosidase (EC 3.2.1.82); GH28 1.51E-04 1.09E-04 4.00E-05 2.65E-05 7.05E-06 2.43E-05 4.90E-05 1.96E-05 7.34E-05 3.78 hemicellulose rhamnogalacturonase (EC 3.2.1.171); rhamnogalacturonan α-1,2- galacturonohydrolase (EC 3.2.1.173); endo- xylogalacturonan hydrolase (EC 3.2.1.-) α,α-trehalase (EC 3.2.1.28); maltose phosphorylase (EC 2.4.1.8); trehalose phosphorylase (EC 2.4.1.64); kojibiose phosphorylase (EC 2.4.1.230); trehalose-6- phosphate phosphorylase (EC 2.4.1.216); nigerose phosphorylase (EC 2.4.1.279); 3-O- GH65 1.36E-04 1.27E-04 5.12E-05 1.63E-05 3.82E-06 4.45E-05 1.19E-05 4.85E-05 6.75E-05 3.78 NA α-glucopyranosyl-L-rhamnose phosphorylase (EC 2.4.1.282); 2-O-α- glucopyranosylglycerol: phosphate β- glucosyltransferase (EC 2.4.1.-); α-glucosyl- 1,2-β-galactosyl-L-hydroxylysine α- glucosidase (EC 3.2.1.107) endoglucanase (EC 3.2.1.4); endo-β-1,4- xylanase (EC 3.2.1.8); β-xylosidase (EC GH51 1.46E-04 1.74E-04 9.83E-05 7.43E-05 7.56E-06 4.17E-05 2.46E-05 4.30E-05 9.52E-06 3.74 NA 3.2.1.37); α-L-arabinofuranosidase (EC 3.2.1.55) β-L-arabinofuranosidase (EC 3.2.1.185); 3-C- GH127 1.05E-04 1.27E-04 3.48E-05 2.30E-05 1.21E-05 3.77E-05 4.55E-05 2.64E-05 3.71E-05 3.74 NA carboxy-5-deoxy-L-xylose (aceric acid) hydrolase α-L-fucosidase (EC 3.2.1.51); α-1,3/1,4-L- GH29 2.12E-04 1.52E-04 5.28E-05 5.34E-05 3.28E-05 6.23E-05 8.35E-05 2.47E-05 6.60E-05 3.39 NA fucosidase (EC 3.2.1.111) 38 bioRxiv preprint doi: https://doi.org/10.1101/440172; this version posted October 11, 2018. The copyright holder for this preprint (which was not certified by peer review) is the author/funder, who has granted bioRxiv a license to display the preprint in perpetuity. It is made available under aCC-BY 4.0 International license.

α-galactosidase (EC 3.2.1.22); α-N- acetylgalactosaminidase (EC 3.2.1.49); isomalto-dextranase (EC 3.2.1.94); β-L- GH27 3.26E-05 1.60E-05 1.76E-05 1.71E-05 1.53E-06 1.22E-06 2.71E-06 4.16E-06 6.75E-06 3.33 galactose arabinopyranosidase (EC 3.2.1.88); galactan:galactan galactosyltransferase (EC 2.4.1.-) chitinase (EC 3.2.1.14); lysozyme (EC GH19 1.98E-05 1.58E-05 5.54E-06 9.38E-06 5.40E-06 5.31E-06 2.42E-06 9.46E-06 0.00E+00 3.32 chitin 3.2.1.17) α-glucosidase (EC 3.2.1.20); α-galactosidase (EC 3.2.1.22); α-mannosidase (EC 3.2.1.24); α-1,3-glucosidase (EC 3.2.1.84); sucrase- isomaltase (EC 3.2.1.48) (EC 3.2.1.10); α- GH31 2.00E-04 1.96E-04 1.08E-04 7.27E-05 1.82E-05 5.43E-05 3.17E-05 3.72E-05 1.48E-04 2.95 α-glucan/mannan xylosidase (EC 3.2.1.177); α-glucan lyase (EC 4.2.2.13); isomaltosyltransferase (EC 2.4.1.-); oligosaccharide α-1,4-glucosyltransferase (EC 2.4.1.161); sulfoquinovosidase (EC 3.2.1.-) GH76 4.69E-05 2.35E-05 8.15E-06 3.42E-05 4.97E-06 2.03E-05 1.07E-05 1.75E-06 6.14E-06 2.86 mannan α-1,6-mannanase (EC 3.2.1.101) xyloglucan:xyloglucosyltransferase (EC 2.4.1.207); keratan-sulfate endo-1,4-β- galactosidase (EC 3.2.1.103); endo-1,3-β- glucanase (EC 3.2.1.39); endo-1,3(4)-β- glucanase (EC 3.2.1.6); licheninase (EC 3.2.1.73); β-agarase (EC 3.2.1.81); κ- β-glucan/ GH16 1.42E-04 1.23E-04 1.20E-04 4.78E-05 2.18E-05 2.20E-05 5.52E-05 1.79E-05 4.30E-05 2.84 carrageenase (EC 3.2.1.83); xyloglucanase hemicellulose (EC 3.2.1.151); endo-β-1,3-galactanase (EC 3.2.1.181); β-porphyranase (EC 3.2.1.178); hyaluronidase (EC 3.2.1.35); endo-β-1,4- galactosidase (EC 3.2.1.-); chitin β-1,6- glucanosyltransferase (EC 2.4.1.-); endo-β- 1,4-galactosidase (EC 3.2.1.-) chitinase (EC 3.2.1.14); lysozyme (EC 3.2.1.17); endo-β-N-acetylglucosaminidase (EC 3.2.1.96); peptidoglycan hydrolase with GH18 2.99E-04 1.00E-04 1.44E-04 1.66E-04 1.02E-05 2.78E-05 3.09E-05 6.04E-05 6.11E-05 2.79 chitin endo-β-N-acetylglucosaminidase specificity (EC 3.2.1.-); Nod factor hydrolase (EC 3.2.1.- ); xylanase inhibitor; concanavalin B; narbonin GH82 5.08E-06 3.43E-07 2.31E-06 0.00E+00 5.17E-07 2.28E-06 2.16E-06 9.89E-08 0.00E+00 2.58 NA Ι-carrageenase (EC 3.2.1.157) GH106 4.64E-05 5.71E-05 4.04E-05 2.17E-05 5.60E-06 1.55E-05 4.12E-05 1.81E-05 0.00E+00 2.54 NA α-L-rhamnosidase (EC 3.2.1.40) GH50 2.06E-05 6.87E-06 7.31E-06 1.73E-06 3.78E-06 1.27E-05 1.36E-05 3.19E-07 0.00E+00 2.44 NA β-agarase (EC 3.2.1.81) peptidoglycan lytic transglycosylase (EC GH102 2.00E-06 1.57E-05 6.45E-07 8.28E-07 6.64E-06 5.24E-06 7.21E-06 1.39E-07 6.14E-06 2.31 NA 3.2.1.-) GH20 2.51E-04 1.04E-04 1.50E-04 1.52E-04 1.02E-05 1.60E-05 5.22E-05 6.07E-05 9.85E-05 2.30 NA β-hexosaminidase (EC 3.2.1.52); lacto-N- 39 bioRxiv preprint doi: https://doi.org/10.1101/440172; this version posted October 11, 2018. The copyright holder for this preprint (which was not certified by peer review) is the author/funder, who has granted bioRxiv a license to display the preprint in perpetuity. It is made available under aCC-BY 4.0 International license.

biosidase (EC 3.2.1.140); β-1,6-N- acetylglucosaminidase) (EC 3.2.1.-); β-6-SO3- N-acetylglucosaminidase (EC 3.2.1.-) α-1,3-L-neoagarooligosaccharide hydrolase GH117 1.58E-05 3.82E-06 9.70E-06 1.90E-06 2.11E-06 6.89E-06 7.28E-06 2.36E-06 0.00E+00 2.27 NA (EC 3.2.1.-); α-1,3-L-neoagarobiase / neoagarobiose hydrolase (EC 3.2.1.-) sialidase or neuraminidase (EC 3.2.1.18); trans-sialidase (EC 2.4.1.-); 2-keto-3- deoxynononic acid hydrolase (EC 3.2.1.-); GH33 5.69E-05 5.64E-05 5.08E-05 2.00E-05 1.47E-05 1.65E-05 3.75E-05 1.66E-05 1.93E-05 2.26 NA anhydrosialidase (EC 4.2.2.15); 3-deoxy-D- manno-octulosonic-acid hydrolase (EC 3.2.1.- ) α-glucuronidase (EC 3.2.1.139); xylan α-1,2- GH67 4.24E-05 6.59E-05 6.70E-05 4.43E-05 1.96E-06 9.16E-06 5.68E-06 3.99E-05 0.00E+00 2.25 NA glucuronidase (EC 3.2.1.131) α-galactosidase (EC 3.2.1.22); α-N- acetylgalactosaminidase (EC 3.2.1.49); GH36 4.98E-05 5.80E-05 2.07E-05 4.08E-05 7.59E-06 2.16E-05 3.17E-05 2.65E-05 2.76E-05 2.14 galactose stachyose synthase (EC 2.4.1.67); raffinose synthase (EC 2.4.1.82) lysozyme (EC 3.2.1.17); mannosylglycoprotein endo-β-N-acetylglucosaminidase GH73 2.20E-04 1.47E-04 1.92E-04 1.52E-04 2.27E-05 4.25E-05 6.24E-05 7.67E-05 9.79E-05 1.99 chitin (EC 3.2.1.96); peptidoglycan hydrolase with endo-β-N-acetylglucosaminidase specificity (EC 3.2.1.-) GH109 9.59E-04 8.62E-04 4.66E-04 3.01E-04 2.55E-04 6.84E-04 8.36E-04 2.92E-04 4.08E-04 1.97 galactose α-N-acetylgalactosaminidase (EC 3.2.1.49) β-xylosidase (EC 3.2.1.37); α-L- arabinofuranosidase (EC 3.2.1.55); arabinanase (EC 3.2.1.99); xylanase (EC 3.2.1.8); galactan 1,3-β-galactosidase (EC GH43 6.34E-04 5.57E-04 5.48E-04 4.73E-04 3.63E-05 1.59E-04 1.09E-04 2.74E-04 5.49E-04 1.94 hemicellulose 3.2.1.145); α-1,2-L-arabinofuranosidase (EC 3.2.1.-); exo-α-1,5-L-arabinofuranosidase (EC 3.2.1.-); [inverting] exo-α-1,5-L-arabinanase (EC 3.2.1.-); β-1,3-xylosidase (EC 3.2.1.-) lysozyme type G (EC 3.2.1.17); peptidoglycan lyase (EC 4.2.2.n1) also known in the GH23 6.00E-04 5.81E-04 4.89E-04 5.61E-04 8.52E-05 1.72E-04 2.03E-04 2.38E-04 4.01E-04 1.92 chitin literature as peptidoglycan lytic transglycosylase; chitinase (EC 3.2.1.14) peptidoglycan lytic transglycosylase (EC GH103 9.03E-06 2.23E-05 3.56E-06 4.81E-06 7.96E-06 1.01E-05 1.60E-05 1.47E-06 1.75E-05 1.79 NA 3.2.1.-) glucoamylase (EC 3.2.1.3); glucodextranase GH15 2.44E-06 4.32E-06 1.30E-06 4.26E-07 1.79E-06 2.28E-06 8.27E-06 1.50E-07 0.00E+00 1.66 NA (EC 3.2.1.70); α,α-trehalase (EC 3.2.1.28); dextran dextrinase (EC 2.4.1.2) β-glucosidase (EC 3.2.1.21); β-galactosidase cellulose/hemicellulos(EC 3.2.1.23); β-mannosidase (EC 3.2.1.25); GH1 9.10E-05 1.09E-04 1.17E-04 9.66E-05 3.68E-06 3.58E-05 2.73E-05 6.79E-05 8.04E-05 1.64 e β-glucuronidase (EC 3.2.1.31); β-xylosidase (EC 3.2.1.37); β-D-fucosidase (EC 3.2.1.38); 40 bioRxiv preprint doi: https://doi.org/10.1101/440172; this version posted October 11, 2018. The copyright holder for this preprint (which was not certified by peer review) is the author/funder, who has granted bioRxiv a license to display the preprint in perpetuity. It is made available under aCC-BY 4.0 International license.

phlorizin hydrolase (EC 3.2.1.62); exo-β-1,4- glucanase (EC 3.2.1.74); 6-phospho-β- galactosidase (EC 3.2.1.85); 6-phospho-β- glucosidase (EC 3.2.1.86); strictosidine β- glucosidase (EC 3.2.1.105); lactase (EC 3.2.1.108); amygdalin β-glucosidase (EC 3.2.1.117); prunasin β-glucosidase (EC 3.2.1.118); vicianin hydrolase (EC 3.2.1.119); raucaffricine β-glucosidase (EC 3.2.1.125); thioglucosidase (EC 3.2.1.147); β- primeverosidase (EC 3.2.1.149); isoflavonoid 7-O-β-apiosyl-β-glucosidase (EC 3.2.1.161); ABA-specific β-glucosidase (EC 3.2.1.175); DIMBOA β-glucosidase (EC 3.2.1.182); β- glycosidase (EC 3.2.1.-); hydroxyisourate hydrolase (EC 3.-.-.-) endoglucanase (EC 3.2.1.4); endo-β-1,3(4)- glucanase / lichenase-laminarinase (EC 3.2.1.6); β-glucosidase (EC 3.2.1.21); lichenase / endo-β-1,3-1,4-glucanase (EC cellulose/hemicellulos3.2.1.73); exo-β-1,4-glucanase / GH9 1.90E-04 1.30E-04 2.92E-04 2.58E-04 1.21E-05 2.37E-05 2.85E-05 9.45E-05 3.71E-05 1.50 e cellodextrinase (EC 3.2.1.74); cellobiohydrolase (EC 3.2.1.91); xyloglucan- specific endo-β-1,4-glucanase / endo- xyloglucanase (EC 3.2.1.151); exo-β- glucosaminidase (EC 3.2.1.165) α-amylase (EC 3.2.1.1); pullulanase (EC 3.2.1.41); cyclomaltodextrin glucanotransferase (EC 2.4.1.19); cyclomaltodextrinase (EC 3.2.1.54); trehalose- 6-phosphate hydrolase (EC 3.2.1.93); oligo-α- glucosidase (EC 3.2.1.10); maltogenic amylase (EC 3.2.1.133); neopullulanase (EC 3.2.1.135); α-glucosidase (EC 3.2.1.20); maltotetraose-forming α-amylase (EC GH13 6.08E-04 5.93E-04 6.69E-04 8.12E-04 5.47E-05 2.83E-04 2.41E-04 3.35E-04 4.00E-04 1.50 NA 3.2.1.60); isoamylase (EC 3.2.1.68); glucodextranase (EC 3.2.1.70); maltohexaose- forming α-amylase (EC 3.2.1.98); maltotriose- forming α-amylase (EC 3.2.1.116); branching enzyme (EC 2.4.1.18); trehalose synthase (EC 5.4.99.16); 4-α-glucanotransferase (EC 2.4.1.25); maltopentaose-forming α-amylase (EC 3.2.1.-) ; amylosucrase (EC 2.4.1.4) ; sucrose phosphorylase (EC 2.4.1.7); malto- oligosyltrehalose trehalohydrolase (EC

41 bioRxiv preprint doi: https://doi.org/10.1101/440172; this version posted October 11, 2018. The copyright holder for this preprint (which was not certified by peer review) is the author/funder, who has granted bioRxiv a license to display the preprint in perpetuity. It is made available under aCC-BY 4.0 International license.

3.2.1.141); isomaltulose synthase (EC 5.4.99.11); malto-oligosyltrehalose synthase (EC 5.4.99.15); amylo-α-1,6-glucosidase (EC 3.2.1.33); α-1,4-glucan: phosphate α- maltosyltransferase (EC 2.4.99.16); 6â€²-P- sucrose phosphorylase (EC 2.4.1.-); amino acid transporter β-glucosidase (EC 3.2.1.21); xylan 1,4-β- xylosidase (EC 3.2.1.37); β- glucosylceramidase (EC 3.2.1.45); β-N- acetylhexosaminidase (EC 3.2.1.52); α-L- arabinofuranosidase (EC 3.2.1.55); glucan 1,3- hemicellulose/β- β-glucosidase (EC 3.2.1.58); glucan 1,4-β- GH3 5.62E-04 3.87E-04 6.55E-04 6.06E-04 3.58E-05 1.68E-04 1.96E-04 2.19E-04 5.11E-04 1.39 glucan glucosidase (EC 3.2.1.74); isoprimeverose- producing oligoxyloglucan hydrolase (EC 3.2.1.120); coniferin β-glucosidase (EC 3.2.1.126); exo-1,3-1,4-glucanase (EC 3.2.1.- ); β-N-acetylglucosaminide phosphorylases (EC 2.4.1.-) xylan α-1,2-glucuronidase (3.2.1.131); α-(4- GH115 4.53E-05 4.44E-05 9.62E-05 6.48E-05 3.03E-07 6.42E-06 3.32E-06 5.78E-05 0.00E+00 1.37 NA O-methyl)-glucuronidase (3.2.1.-) endo-β-1,4-xylanase (EC 3.2.1.8); β- glucosidase (3.2.1.21); β-glucuronidase (EC 3.2.1.31); β-xylosidase (EC 3.2.1.37); β- fucosidase (EC 3.2.1.38); glucosylceramidase GH30 1.21E-04 1.14E-04 2.65E-04 1.77E-04 2.61E-06 1.37E-05 1.66E-05 1.06E-04 6.17E-05 1.28 NA (EC 3.2.1.45); β-1,6-glucanase (EC 3.2.1.75); glucuronoarabinoxylan endo-β-1,4-xylanase (EC 3.2.1.136); endo-β-1,6-galactanase (EC:3.2.1.164); [reducing end] β-xylosidase (EC 3.2.1.-) α-amylase (EC 3.2.1.1); α-galactosidase (EC 3.2.1.22); amylopullulanase (EC 3.2.1.41); GH57 1.77E-04 1.84E-04 3.09E-04 3.57E-04 2.37E-05 4.53E-05 5.92E-05 1.47E-04 1.04E-04 1.21 NA cyclomaltodextrinase (EC 3.2.1.54); branching enzyme (EC 2.4.1.18); 4-α-glucanotransferase (EC 2.4.1.25) β-galactosidase (EC 3.2.1.23); α-L- GH42 4.56E-05 4.89E-05 7.39E-05 9.30E-05 5.92E-06 3.44E-05 1.27E-05 2.71E-05 2.76E-05 1.20 NA arabinopyranosidase (EC 3.2.1.-) β-glucosidase (EC 3.2.1.21); β-xylosidase (EC 3.2.1.37); acid β-glucosidase/β- GH116 1.68E-05 1.46E-05 2.09E-05 3.54E-05 3.38E-06 6.71E-06 1.87E-05 6.10E-06 0.00E+00 1.20 NA glucosylceramidase (EC 3.2.1.45); β-N- acetylglucosaminidase (EC 3.2.1.52) amylomaltase or 4-α-glucanotransferase (EC GH77 1.00E-04 1.54E-04 2.42E-04 1.97E-04 1.13E-05 6.10E-05 6.12E-05 9.12E-05 1.08E-04 1.16 NA 2.4.1.25) GH37 3.33E-05 3.86E-05 7.09E-05 8.93E-05 3.83E-07 8.56E-06 8.05E-06 4.89E-05 2.43E-05 1.00 NA α,α-trehalase (EC 3.2.1.28)

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β-1,4-mannosylglucose phosphorylase (EC 2.4.1.281); β-1,4-mannooligosaccharide phosphorylase (EC 2.4.1.319); β-1,4- mannosyl-N-acetyl-glucosamine GH130 7.02E-05 6.18E-05 1.27E-04 1.25E-04 5.02E-06 4.21E-05 3.40E-05 5.85E-05 9.12E-05 0.96 NA phosphorylase (EC 2.4.1.320); β-1,2- mannobiose phosphorylase (EC 2.4.1.-); β- 1,2-oligomannan phosphorylase (EC 2.4.1.-); β-1,2-mannosidase (EC 3.2.1.-) endoglucanase (EC 3.2.1.4); oligoxyloglucan cellulose/hemicellulos GH74 5.17E-05 1.36E-04 3.40E-04 1.21E-04 6.89E-05 3.54E-05 8.05E-05 2.75E-05 5.59E-05 0.90 reducing end-specific cellobiohydrolase (EC e 3.2.1.150); xyloglucanase (EC 3.2.1.151) cellobiose phosphorylase (EC 2.4.1.20); laminaribiose phosphorylase (EC 2.4.1.31); cellodextrin phosphorylase (EC 2.4.1.49); GH94 6.02E-05 1.25E-04 2.30E-04 2.81E-04 4.01E-06 3.49E-05 2.18E-05 1.37E-04 1.04E-05 0.90 cellulose chitobiose phosphorylase (EC 2.4.1.-); cyclic β-1,2-glucan synthase (EC 2.4.1.-); cellobionic acid phosphorylase (EC 2.4.1.321); β-1,2- oligoglucan phosphorylase (EC 2.4.1.-) GH120 1.78E-05 2.38E-05 1.43E-05 4.07E-05 1.40E-05 2.15E-05 2.43E-05 5.02E-06 4.42E-05 0.89 hemicellulose β-xylosidase (EC 3.2.1.37) maltose-6-phosphate glucosidase (EC 3.2.1.122); α-glucosidase (EC 3.2.1.20); α- galactosidase (EC 3.2.1.22); 6-phospho-β- GH4 1.09E-04 9.76E-05 1.98E-04 1.54E-04 2.93E-05 1.32E-04 9.60E-05 8.95E-05 1.36E-04 0.87 NA glucosidase (EC 3.2.1.86); α-glucuronidase (EC 3.2.1.139); α-galacturonase (EC 3.2.1.67); palatinase (EC 3.2.1.-) chitosanase (EC 3.2.1.132); cellulase (EC 3.2.1.4); licheninase (EC 3.2.1.73); endo-1,4- GH8 5.38E-05 7.10E-05 1.76E-04 2.21E-04 6.45E-06 9.81E-06 4.16E-06 5.89E-05 5.28E-05 0.82 cellulose β-xylanase (EC 3.2.1.8); reducing-end-xylose releasing exo-oligoxylanase (EC 3.2.1.156) α-mannosidase (EC 3.2.1.24); mannosyl- oligosaccharide α-1,2-mannosidase (EC 3.2.1.113); mannosyl-oligosaccharide α-1,3- GH38 3.70E-05 4.10E-05 6.96E-05 3.04E-05 2.89E-05 6.31E-05 8.18E-05 3.03E-05 4.33E-05 0.79 mannan 1,6-mannosidase (EC 3.2.1.114); α-2-O- mannosylglycerate hydrolase (EC 3.2.1.170); mannosyl-oligosaccharide α-1,3-mannosidase (EC 3.2.1.-) GH53 2.80E-05 2.08E-05 6.45E-05 1.05E-04 1.77E-06 1.33E-05 6.71E-06 3.29E-05 2.21E-05 0.69 NA endo-β-1,4-galactanase (EC 3.2.1.89) α-L-iduronidase (EC 3.2.1.76); β-xylosidase GH39 5.14E-05 4.21E-05 1.86E-04 1.19E-04 7.72E-06 2.82E-05 3.29E-05 7.36E-05 7.03E-05 0.63 hemicellulose (EC 3.2.1.37) β-mannanase (EC 3.2.1.78); exo-β-1,4- mannobiohydrolase (EC 3.2.1.100); β-1,3- GH26 3.95E-05 3.75E-05 1.53E-04 1.20E-04 1.22E-05 1.94E-05 2.46E-05 4.67E-05 5.25E-05 0.63 hemicellulose xylanase (EC 3.2.1.32); lichenase / endo-β- 1,3-1,4-glucanase (EC 3.2.1.73); mannobiose- producing exo-β-mannanase (EC 3.2.1.-)

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endo-1,4-β-xylanase (EC 3.2.1.8); endo-1,3-β- GH10 1.42E-04 2.07E-04 5.74E-04 6.14E-04 1.41E-05 5.18E-05 5.61E-05 3.87E-04 3.09E-04 0.61 hemicellulose xylanase (EC 3.2.1.32); tomatinase (EC 3.2.1.- ); xylan endotransglycosylase (EC 2.4.2.-) endo-β-1,4-glucanase / cellulase (EC 3.2.1.4); endo-β-1,4-xylanase (EC 3.2.1.8); β- glucosidase (EC 3.2.1.21); β-mannosidase (EC 3.2.1.25); β-glucosylceramidase (EC 3.2.1.45); glucan β-1,3-glucosidase (EC 3.2.1.58); licheninase (EC 3.2.1.73); exo-β- 1,4-glucanase / cellodextrinase (EC 3.2.1.74); glucan endo-1,6-β-glucosidase (EC 3.2.1.75); mannan endo-β-1,4-mannosidase (EC cellulose/hemicellulos3.2.1.78); cellulose β-1,4-cellobiosidase (EC GH5 2.56E-04 2.72E-04 1.03E-03 1.07E-03 5.29E-05 1.32E-04 1.56E-04 4.23E-04 3.42E-04 0.58 e 3.2.1.91); steryl β-glucosidase (EC 3.2.1.104); endoglycoceramidase (EC 3.2.1.123); chitosanase (EC 3.2.1.132); β-primeverosidase (EC 3.2.1.149); xyloglucan-specific endo-β- 1,4-glucanase (EC 3.2.1.151); endo-β-1,6- galactanase (EC 3.2.1.164); hesperidin 6-O-α- L-rhamnosyl-β-glucosidase (EC 3.2.1.168); β- 1,3-mannanase (EC 3.2.1.-); arabinoxylan- specific endo-β-1,4-xylanase (EC 3.2.1.-); mannan transglycosylase (EC 2.4.1.-) endo-β-1,4-xylanase (EC 3.2.1.8); endo-β-1,3- GH11 3.03E-05 5.94E-05 2.76E-04 3.97E-04 3.33E-06 1.24E-05 1.97E-05 1.66E-04 2.16E-04 0.29 hemicellulose xylanase (EC 3.2.1.32) GH129 2.64E-06 4.26E-06 2.71E-05 3.84E-05 1.22E-06 1.27E-05 7.13E-06 6.50E-06 0.00E+00 0.26 NA α-N-acetylgalactosaminidase (EC 3.2.1.49) GH113 2.20E-06 2.75E-06 5.48E-06 3.68E-05 1.31E-06 6.24E-06 4.16E-06 4.25E-06 1.44E-05 0.24 hemicellulose β-mannanase (EC 3.2.1.78) cellulose/hemicellulosendoglucanase (EC 3.2.1.4); xyloglucanase GH44 2.57E-06 5.03E-08 1.72E-05 2.08E-06 2.30E-06 5.26E-06 7.95E-07 2.35E-06 8.60E-06 0.24 e (EC 3.2.1.151) cellulose/hemicellulos GH45 5.40E-07 1.18E-05 7.98E-05 1.88E-04 8.70E-08 6.87E-07 1.74E-07 1.97E-05 3.35E-05 0.13 endoglucanase (EC 3.2.1.4) e GH46 0.00E+00 0.00E+00 1.28E-07 2.95E-07 1.21E-07 0.00E+00 3.46E-07 0.00E+00 0.00E+00 0.00 NA chitosanase (EC 3.2.1.132) GH52 0.00E+00 0.00E+00 1.43E-05 2.68E-06 0.00E+00 2.55E-07 1.39E-07 6.91E-06 0.00E+00 0.00 hemicellulose β-xylosidase (EC 3.2.1.37) α-L-arabinofuranosidase (EC 3.2.1.55); β- GH54 0.00E+00 0.00E+00 1.19E-07 0.00E+00 1.93E-06 3.57E-07 1.29E-06 0.00E+00 0.00E+00 0.00 hemicellulose xylosidase (EC 3.2.1.37). GH62 0.00E+00 0.00E+00 1.21E-06 0.00E+00 1.88E-06 1.32E-06 6.09E-07 4.54E-07 0.00E+00 0.00 hemicellulose α-L-arabinofuranosidase (EC 3.2.1.55) GH71 0.00E+00 0.00E+00 0.00E+00 0.00E+00 0.00E+00 3.65E-06 1.25E-06 1.51E-07 0.00E+00 0.00 β-glucan α-1,3-glucanase (EC 3.2.1.59) GH75 0.00E+00 0.00E+00 0.00E+00 0.00E+00 1.54E-07 2.72E-07 3.99E-07 0.00E+00 0.00E+00 0.00 NA chitosanase (EC 3.2.1.132) β-glucuronidase (EC 3.2.1.31); hyaluronoglucuronidase (EC 3.2.1.36); GH79 0.00E+00 0.00E+00 9.32E-06 8.72E-06 0.00E+00 1.01E-06 8.73E-07 5.70E-07 6.45E-06 0.00 NA heparanase (EC 3.2.1.166); baicalin β- glucuronidase (EC 3.2.1.167); β-4-O-methyl- 44 bioRxiv preprint doi: https://doi.org/10.1101/440172; this version posted October 11, 2018. The copyright holder for this preprint (which was not certified by peer review) is the author/funder, who has granted bioRxiv a license to display the preprint in perpetuity. It is made available under aCC-BY 4.0 International license.

glucuronidase (EC 3.2.1.-) N-acetyl β-glucosaminidase (EC 3.2.1.52); hyaluronidase (EC 3.2.1.35); [protein]-3-O- GH84 0.00E+00 0.00E+00 1.26E-06 1.22E-06 3.89E-07 0.00E+00 2.93E-06 3.50E-07 0.00E+00 0.00 NA (GlcNAc)-L-Ser/Thr β-N- acetylglucosaminidase (EC 3.2.1.169) inulin lyase [DFA-I-forming] (EC 4.2.2.17); inulin lyase [DFA-III-forming] (EC 4.2.2.18); GH91 0.00E+00 0.00E+00 0.00E+00 3.64E-07 8.90E-08 2.63E-07 4.34E-07 0.00E+00 0.00E+00 0.00 NA difructofuranose 1,2':2,3' dianhydride hydrolase [DFA-IIIase] (EC 3.2.1.-)

45 bioRxiv preprint doi: https://doi.org/10.1101/440172; this version posted October 11, 2018. The copyright holder for this preprint (which was not certified by peer review) is the author/funder, who has granted bioRxiv a license to display the preprint in perpetuity. It is made available under aCC-BY 4.0 International license.

Table S3. Relative abundance of plant-degrading enzymes in gut metagenomes of nine termites with different diets. Mn: Macrotermes natalensis, Od: Odontotermes sp., Nc: Nasutitermes corniger, Aw: Amitermes wheeleri, Mp: Microcerotermes parvus, Co: Cornitermes sp., Th: Termes hospes, Nt: Neocapritermes taracua, Cu: Cubitermes ugandensis.

Substrate Enzyme activity EC # Mn Od Cu Aw Nt Th Co Mp Nc beta-1,4-Endoglucanase 3.2.1.4 1.91E-04 2.31E-04 2.79E-05 2.87E-04 6.85E-05 6.36E-05 3.91E-04 1.01E-03 1.29E-03 Cellobiohydrolase (reducing end) 3.2.1.176 0.00E+00 0.00E+00 1.39E-07 0.00E+00 0.00E+00 0.00E+00 0.00E+00 0.00E+00 0.00E+00 Cellobiohydrolase (nonreducing end) 3.2.1.91 7.34E-06 6.90E-06 3.75E-07 0.00E+00 1.19E-06 2.38E-06 2.54E-05 1.83E-05 2.85E-05 Cellulose beta-1,4-Glucosidase 3.2.1.21 4.16E-04 2.51E-04 5.47E-06 1.20E-04 3.97E-05 4.75E-05 9.58E-05 2.50E-04 2.07E-04 Cellobiose dehydrogenase 1.1.99.18 0.00E+00 0.00E+00 0.00E+00 0.00E+00 0.00E+00 0.00E+00 0.00E+00 0.00E+00 0.00E+00 Pyranose dehydrogenase 1.1.99.29 0.00E+00 0.00E+00 0.00E+00 0.00E+00 0.00E+00 0.00E+00 0.00E+00 0.00E+00 0.00E+00 Lytic polysaccharide monooxygenase LPMO 0.00E+00 0.00E+00 1.83E-07 0.00E+00 0.00E+00 0.00E+00 0.00E+00 0.00E+00 0.00E+00 beta-1,4-Endomannanase 3.2.1.78 2.40E-04 3.68E-04 1.62E-05 4.36E-04 7.42E-05 7.21E-05 4.97E-04 7.52E-04 9.48E-04 beta-1,4-Mannosidase 3.2.1.25 1.89E-04 1.80E-04 2.92E-06 4.21E-05 2.14E-05 3.86E-05 8.50E-05 1.46E-04 8.42E-05 Galactomannan beta-1,4-Galactosidase 3.2.1.23 5.14E-05 4.87E-05 1.20E-05 4.33E-05 2.07E-05 2.19E-05 5.60E-05 1.79E-04 1.76E-04 alpha-1,4-Galactosidase 3.2.1.22 3.19E-05 2.11E-05 2.32E-06 0.00E+00 1.28E-05 3.79E-05 1.21E-05 4.50E-05 3.61E-05 alpha-Arabinofuranosidase 3.2.1.55 5.36E-04 4.01E-04 1.42E-05 2.52E-05 6.62E-05 7.63E-05 9.81E-05 1.83E-04 1.86E-04 Xyloglucan beta-1,4-endoglucanase 3.2.1.151 2.29E-04 1.78E-04 1.77E-05 9.55E-05 6.76E-05 6.48E-05 6.23E-05 1.15E-04 1.31E-04 alpha-Arabinofuranosidase 3.2.1.55 5.26E-04 5.11E-04 1.62E-05 3.13E-04 6.42E-05 1.08E-04 2.80E-04 5.75E-04 4.93E-04 alpha-Xylosidase 3.2.1.177 3.83E-05 3.21E-05 2.23E-06 0.00E+00 5.03E-06 3.68E-06 2.22E-05 7.32E-05 6.60E-05 Xyloglucan alpha-Fucosidase 3.2.1.51 5.26E-04 5.11E-04 1.62E-05 3.13E-04 6.42E-05 1.08E-04 2.80E-04 5.75E-04 4.93E-04 alpha-1,4-Galactosidase 3.2.1.22 1.61E-04 1.56E-04 3.37E-06 2.00E-05 7.13E-06 4.07E-05 3.01E-05 7.03E-05 5.82E-05 beta-1,4-Galactosidase 3.2.1.23 3.29E-04 2.62E-04 2.37E-05 3.28E-05 7.43E-05 6.12E-05 2.68E-05 7.58E-05 6.34E-05 Xylan_alpha-1,2-glucuronosidase 3.2.1.131 2.29E-04 1.78E-04 1.77E-05 9.55E-05 6.76E-05 6.48E-05 6.23E-05 1.15E-04 1.31E-04 beta-1,4-Endoxylanase 3.2.1.8 5.36E-04 4.01E-04 1.42E-05 2.52E-05 6.62E-05 7.63E-05 9.81E-05 1.83E-04 1.86E-04 Xylan beta-1,4-Xylosidase 3.2.1.37 4.24E-05 6.59E-05 1.96E-06 0.00E+00 4.48E-06 9.16E-06 3.98E-05 6.70E-05 4.43E-05 Arabinoxylan arabinofuranohydrolase/arabinofuranosidase 3.2.1.55 5.26E-04 5.11E-04 1.62E-05 3.13E-04 6.42E-05 1.08E-04 2.80E-04 5.75E-04 4.93E-04 alpha-Glucuronidase 3.2.1.139 0.00E+00 0.00E+00 0.00E+00 0.00E+00 1.20E-06 0.00E+00 0.00E+00 0.00E+00 0.00E+00 alpha-1,4-Galactosidase 3.2.1.22 2.29E-04 1.78E-04 1.77E-05 9.55E-05 6.76E-05 6.48E-05 6.23E-05 1.15E-04 1.31E-04 beta-1,4-Galactosidase 3.2.1.23 5.36E-04 4.01E-04 1.42E-05 2.52E-05 6.62E-05 7.63E-05 9.81E-05 1.83E-04 1.86E-04 Arabinoxylan Acetyl xylan esterase 3.1.1.72 1.66E-04 2.28E-04 7.05E-06 6.14E-06 3.03E-05 2.70E-05 2.90E-05 1.93E-05 7.05E-05 Feruloyl esterase 3.1.1.73 1.15E-04 1.97E-04 1.92E-06 0.00E+00 1.30E-05 7.63E-06 8.56E-06 3.20E-05 8.71E-05 beta-1,4-Endogalactanase 3.2.1.89 2.63E-05 2.02E-05 1.77E-06 6.75E-06 3.81E-06 1.13E-05 3.08E-05 6.28E-05 1.03E-04 Endoarabinanase 3.2.1.99 3.79E-05 2.43E-05 1.71E-06 2.89E-05 8.40E-06 5.71E-06 6.30E-06 2.86E-05 6.24E-06 Galactan_1,3-beta-galactosidase 3.2.1.145 0.00E+00 0.00E+00 0.00E+00 0.00E+00 0.00E+00 0.00E+00 0.00E+00 0.00E+00 0.00E+00 Acetylesterase 3.1.1.6 0.00E+00 0.00E+00 0.00E+00 0.00E+00 0.00E+00 0.00E+00 0.00E+00 0.00E+00 0.00E+00 Endopolygalacturonases 3.2.1.15 9.21E-05 6.10E-05 3.02E-07 3.87E-05 9.17E-06 6.50E-06 1.16E-05 2.06E-05 2.04E-05 Pectin Exopolygalacturonases 3.2.1.67 1.04E-05 1.42E-05 3.90E-07 7.06E-06 3.43E-06 1.72E-05 1.59E-05 5.10E-05 2.57E-05 Endorhamnogalacturonase 3.2.1.171 0.00E+00 0.00E+00 0.00E+00 0.00E+00 0.00E+00 0.00E+00 0.00E+00 0.00E+00 0.00E+00 46 bioRxiv preprint doi: https://doi.org/10.1101/440172; this version posted October 11, 2018. The copyright holder for this preprint (which was not certified by peer review) is the author/funder, who has granted bioRxiv a license to display the preprint in perpetuity. It is made available under aCC-BY 4.0 International license.

Rhamnogalacturonan rhamnohydrolase 3.2.1.174 0.00E+00 0.00E+00 0.00E+00 0.00E+00 0.00E+00 0.00E+00 0.00E+00 0.00E+00 0.00E+00 alpha-Rhamnosidase 3.2.1.40 0.00E+00 0.00E+00 0.00E+00 0.00E+00 0.00E+00 0.00E+00 0.00E+00 0.00E+00 0.00E+00 alpha-Arabinofuranosidase 3.2.1.55 5.26E-04 5.11E-04 1.62E-05 3.13E-04 6.42E-05 1.08E-04 2.80E-04 5.75E-04 4.93E-04 Exoarabinanase 3.2.1.- 1.44E-05 1.29E-05 4.50E-07 0.00E+00 3.67E-06 2.55E-06 2.59E-06 3.54E-06 3.89E-06 Unsaturated glucuronyl hydrolase 3.2.1.- 1.62E-06 9.73E-06 8.65E-08 6.14E-06 3.12E-06 4.43E-06 2.77E-06 3.13E-06 3.17E-07 Unsaturated rhamnogalacturonan hydrolase 3.2.1.172 1.40E-04 1.10E-04 1.24E-06 4.88E-05 1.36E-05 2.88E-05 2.90E-05 7.37E-05 4.35E-05 beta-1,4-Xylosidase 3.2.1.37 1.89E-04 1.80E-04 2.92E-06 4.21E-05 2.14E-05 3.86E-05 8.50E-05 1.46E-04 8.42E-05 beta-1,4-Galactosidase 3.2.1.23 5.36E-04 4.01E-04 1.42E-05 2.52E-05 6.62E-05 7.63E-05 9.81E-05 1.83E-04 1.86E-04 Pectin lyase 4.2.2.10 7.21E-07 0.00E+00 0.00E+00 0.00E+00 0.00E+00 0.00E+00 8.09E-08 0.00E+00 0.00E+00 Pectate lyase 4.2.2.2 5.19E-05 6.44E-05 0.00E+00 8.60E-06 1.97E-06 4.67E-06 1.29E-05 2.89E-05 8.34E-06 Rhamnogalacturonan lyase 4.2.2.23 2.88E-05 3.58E-05 1.22E-06 0.00E+00 1.14E-05 1.21E-05 1.03E-05 4.03E-05 5.35E-05 Pectin methyl esterase 3.1.1.11 7.26E-05 5.67E-05 0.00E+00 0.00E+00 2.83E-07 1.98E-06 5.31E-06 1.64E-06 0.00E+00 Pectin acetyl esterase/Rhamnogalacturonan acetyl esterase 3.1.1.- 7.10E-05 4.88E-05 6.39E-06 3.99E-05 1.10E-05 8.16E-06 1.98E-06 1.16E-05 2.53E-05 Feruloyl esterase 3.1.1.73 1.15E-04 1.97E-04 1.92E-06 0.00E+00 1.30E-05 7.63E-06 8.56E-06 3.20E-05 8.71E-05 laccase 1.10.3.2 0.00E+00 0.00E+00 0.00E+00 0.00E+00 0.00E+00 0.00E+00 0.00E+00 2.54E-07 5.63E-06 Lignin 4-O-methyl-glucuronoyl methylesterase 3.1.1.- 3.75E-05 5.76E-05 1.09E-05 2.76E-05 3.85E-05 3.16E-05 1.13E-05 2.50E-05 7.83E-05 versatile_peroxidase 1.11.1.16 0.00E+00 0.00E+00 0.00E+00 0.00E+00 0.00E+00 0.00E+00 0.00E+00 0.00E+00 0.00E+00

47 bioRxiv preprint doi: https://doi.org/10.1101/440172; this version posted October 11, 2018. The copyright holder for this preprint (which was not certified by peer review) is the author/funder, who has granted bioRxiv a license to display the preprint in perpetuity. It is made available under aCC-BY 4.0 International license.

Table S4. Relative abundance of the main catalytic types of proteases in the gut metagenomes of nine termite species with different diets. Mn: Macrotermes natalensis, Od: Odontotermes sp., Nc: Nasutitermes corniger, Aw: Amitermes wheeleri, Mp: Microcerotermes parvus, Co: Cornitermes sp., Th: Termes hospes, Nt: Neocapritermes taracua, Cu: Cubitermes ugandensis.

Type of protease Mn Od Cu Aw Nt Th Co Mp Nc Aspartic Peptidases 5.66E-04 5.75E-04 4.36E-04 3.93E-04 3.78E-04 3.94E-04 2.60E-04 4.69E-04 4.59E-04 Cysteine Peptidases 5.03E-03 5.29E-03 1.29E-03 4.86E-03 2.35E-03 2.74E-03 2.81E-03 4.33E-03 4.16E-03 Glutamic Peptidases 1.67E-06 6.38E-06 3.41E-06 1.00E-07 2.32E-06 8.36E-07 1.00E-07 1.00E-07 2.66E-07 Metallo Peptidases 1.09E-02 1.14E-02 2.74E-03 9.83E-03 7.24E-03 6.07E-03 4.94E-03 1.07E-02 1.01E-02 Asparagine Peptide Lyases 8.28E-04 6.91E-04 7.94E-05 4.90E-04 3.12E-04 3.29E-04 2.57E-04 5.64E-04 5.96E-04 Mixed Peptidases 2.00E-05 2.62E-05 6.04E-06 4.91E-06 1.56E-05 8.35E-06 2.92E-06 9.05E-06 1.26E-05 Serine Peptidases 9.39E-03 9.34E-03 2.13E-03 7.57E-03 5.30E-03 4.54E-03 4.16E-03 6.63E-03 6.52E-03 Threonine Peptidases 4.85E-04 2.89E-04 6.51E-04 6.09E-04 1.09E-03 7.34E-04 4.81E-04 4.97E-04 6.56E-04 Peptidases of Unknown Type 1.20E-03 1.21E-03 3.46E-04 9.15E-04 7.93E-04 7.31E-04 5.45E-04 6.97E-04 7.08E-04 Sum 2.85E-02 2.88E-02 7.69E-03 2.47E-02 1.75E-02 1.56E-02 1.35E-02 2.39E-02 2.32E-02