Terrestrial Microbiology

Temporal dynamics of fibrolytic and methanogenic rumen microorganisms during in situ incubation of switchgrass determined by 16S rRNA gene profiling

Hailan Piao, Medora Lachman, Stephanie Malfatti, Alexander Sczyrba, Bernhard Knierim, Manfred Auer, Susannah Green

Tringe, Roderick Ian Mackie, Carl James Yeoman and Matthias Hess

Journal Name: Frontiers in Microbiology

ISSN: 1664-302X

Article type: Original Research Article

Received on: 09 May 2014

Accepted on: 03 Jun 2014

Provisional PDF published on: 03 Jun 2014

www.frontiersin.org: www.frontiersin.org

Citation: Piao H, Lachman M, Malfatti S, Sczyrba A, Knierim B, Auer M, Tringe SG, Mackie RI, Yeoman CJ and Hess M(2014) Temporal dynamics of fibrolytic and methanogenic rumen microorganisms during in situ incubation of switchgrass determined by 16S rRNA gene profiling. Front. Microbiol. 5:307. doi:10.3389/fmicb.2014.00307

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Copyright statement: © 2014 Piao, Lachman, Malfatti, Sczyrba, Knierim, Auer, Tringe, Mackie, Yeoman and Hess. This is an open-access article distributed under the terms of the Creative Commons Attribution License (CC BY). The use, distribution or reproduction in other forums is permitted, provided the original author(s) or licensor are credited and that the original publication in this journal is cited, in accordance with accepted academic practice. No use, distribution or reproduction is permitted which does not comply with these terms.

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1 Temporal dynamics of fibrolytic and methanogenic rumen microorganisms during 2 in situ incubation of switchgrass determined by 16S rRNA gene profiling 3 4 Hailan Piaoa), Medora Lachmanb), Stephanie Malfattic), Alexander Sczyrbad), Bernhard 5 Knierime), Manfred Auere), Susannah Tringef), Roderick I. Mackieg), Carl J. Yeomand), 6 *Matthias Hessa,f,h 7 8 a) Washington State University, Richland, WA, 99354, USA 9 b) Montana State University, Bozeman, MT, 59718, USA 10 c) Lawrence Livermore National Laboratory, Livermore, CA, 94550, USA 11 d) University of Bielefeld, Bielefeld, D-33501, Germany 12 e) Lawrence Berkeley National Laboratory, Berkeley, CA, 94598, USA 13 f) DOE Joint Genome Institute, Walnut Creek, CA, 94598, USA 14 g) University of Illinois, Urbana-Champaign, IL 61801, USA 15 h) Pacific Northwest National Laboratory, Richland, WA, 99352, USA 16 17 * Correspondence: 18 Dr. Matthias Hess 19 Systems Microbiology & Biotechnology Group 20 Washington State University 21 2710 Crimson Way 22 Richland, WA 99354-1671, USA 23 [email protected] 24 25 26 Abstract: 27 The rumen microbial ecosystem is known for its biomass-degrading and methane- 28 producing phenotype. Fermentation of recalcitrant plant material, comprised of a 29 multitude of interwoven fibers, necessitates the synergistic activity of diverse microbial 30 taxonomic groups that inhabit the anaerobic rumen ecosystem. Although interspecies 31 hydrogen (H2) transfer, a process during which bacterially generated H2 is transferred to 32 methanogenic Archaea, has obtained significant attention over the last decades, the 33 temporal variation of the different taxa involved in in situ biomass-degradation, H2 34 transfer and the methanogenesis process remains to be established. Here we investigated 35 the temporal succession of microbial taxa and its effect on fiber composition during 36 rumen incubation using 16S rRNA amplicon sequencing. Switchgrass filled nylon bags 37 were placed in the rumen of a cannulated cow and collected at nine time points for DNA 38 extraction and 16S pyrotag profiling. The microbial community colonizing the air-dried 39 and non-incubated (0 h) switchgrass was dominated by members of the Bacilli (recruiting 40 63 % of the pyrotag reads). During in situ incubation of the switchgrass, two major shifts 41 in the community composition were observed: Bacilli were replaced within 30 min by 42 members belonging to the Bacteroidia and Clostridia, which recruited 34 % and 25 % of 43 the 16S rRNA reads generated, respectively. A second significant shift was observed 44 after 16 h of rumen incubation, when members of the Spirochaetes and Fibrobacteria 45 classes became more abundant in the fiber-adherent community. During the first 30 min 46 of rumen incubation ~13 % of the switchgrass dry matter was degraded, whereas little 47 biomass degradation appeared to have occurred between 30 min and 4 h after the 48 switchgrass was placed in the rumen. Interestingly, methanogenic members of the 49 Euryarchaeota (i.e. Methanobacteria) increased up to 3-fold during this period of 50 reduced biomass-degradation, with peak abundance just before rates of dry matter 51 degradation increased again. We hypothesize that during this period microbial-mediated 52 fibrolysis was temporarily inhibited until H2 was metabolized into CH4 by methanogens. 53 Collectively, our results demonstrate the importance of inter-species interactions for the 54 biomass-degrading and methane-producing phenotype of the rumen microbiome – both 55 microbially facilitated processes with global significance. 56 Keywords: Rumen Microbiology, microbe-microbe interactions, cellulolytic bacteria, 57 methanogenic archaea, interspecies H2 transfer 58 59 Introduction: 60 The microbial community (microbiota) inhabiting the cow rumen has been 61 described as “the most elegant and highly evolved cellulose-digesting system in nature” 62 (Weimer et al, 2009). Cellulose, the most abundant natural polymer on earth (Kadla & 63 Gilbert, 2000), and a major component of plant biomass (Heredia et al, 1995) is degraded 64 within the rumen by various bacteria (Weimer, 1996), fungi (Bauchop & Mountfort, 1981; 65 Theodorou et al, 1996), and protozoa (Coleman, 1992). The efficacy of cellulose- 66 degradation, and ultimately fiber-degradation, is mediated by important microbial 67 enzymatic and metabolic interactions and it has been shown that major cellulolytic 68 bacteria, such as Fibrobacter succinogenes, Ruminococcus flavefaciens, and 69 Ruminococcus albus (Weimer, 1996), exhibit suboptimal cellulolytic activity if they are 70 not able to synergistically engage with non-cellulolytic microorganisms (Fondevila & 71 Dehority, 1996; Kudo et al, 1987). In particular methanogenic archaea have been 72 described to boost the carbohydrate-degrading activity of cellulolytic rumen bacteria 73 (Joblin et al, 1989). The role of rumen methanogens is unique in that these organisms do 74 not degrade or ferment any portion of plant biomass, but instead obtain their energy from 75 byproducts, principally H2 and CO2, of fibrolytic organisms (Janssen & Kirs, 2008). By 76 using hydrogen to reduce CO2, methanogens remove otherwise inhibitory levels of H2 77 (Janssen & Kirs, 2008), increase ATP yields by redirecting reducing equivalents toward 78 acetate (Latham & Wolin, 1977) and thereby promote growth (Rychlik & May, 2000) 79 and the ability to produce higher concentrations of fibrolytic enzymes. Physical co- 80 aggregation between fibrolytic and methanogenic populations are commonly observed 81 [i.e. (Leahy et al, 2010)] and this is thought to be important for both the efficient transfer 82 of H2 and maintaining the low H2 partial pressures necessary to sustain active growth of 83 the fermentative bacteria (Ishii et al, 2006; Stams, 1994). 84 85 To this end, the temporal dynamics of fibrolytic bacteria and methanogenic archaea 86 remain to be systematically explored during in situ colonization and degradation of plant 87 biomass within the rumen. Previous studies have shown temporal changes in bacterial 88 (Edwards et al, 2007; Huws et al, 2013) and fungal (Edwards et al, 2008) populations 89 during ruminal incubation of fresh perennial rye grass (Lolium perenne). Specifically, 90 these studies revealed a rapid colonization of the plant material within 5 min of rumen- 91 incubation by both bacterial (Edwards et al, 2007) and fungal populations, followed by a 92 compositional shift in bacterial populations between 2-4 h following incubation (Huws et 93 al, 2013). Compositional dynamics may vary among forage types based on observed 94 differences in fiber disappearance and fibrolytic enzyme activities (Bowman & Firkins, 95 1993) and the observed temporal changes in microbiota may be affected by residual plant 96 metabolism of fresh forages (Huws et al, 2013). Here we provide evidence that similar 97 temporal changes occur during the in situ incubation of dried switchgrass and correspond 98 to changes in methanogen abundance. Furthermore, these changes in microbiota also 99 correspond to notable differences in the rate of forage degradation. 100 101 Materials and Methods: 102 Sample collection 103 To enrich for fiber-adherent rumen microorganisms, air-dried switchgrass 104 (Panicum virgatum) was ground into 2 mm pieces using a Wiley mill and weighed into 105 individual in situ nylon bags (50 µm pores; Ankom Technology, Macedon, NY, USA). A 106 total of 18 nylon bags, each containing 5 g of air-dried switchgrass, were placed in the 107 rumen of one cannulated Friesian cow, fed on a mixed diet containing 60 % fiber (Hess et 108 al, 2011). Nylon bags were retrieved from the cow’s rumen at 0.5, 1, 2, 4, 6, 16, 24, 48, 109 and 72 h and washed immediately with 2 x 50 ml PBS buffer (pH7) to remove rumen 110 fluid containing loosely adherent microbes, placed on dry-ice and transported 111 immediately to the laboratory where DNA extraction and fiber degradation analysis were 112 performed. Non-incubated nylon bags filled with ground switchgrass were used as 113 control (0 h). To account for biological variation during the fiber colonization process, 114 nylon bags were retrieved from the cow’s rumen in duplicates. All animal procedures 115 were carried out under an approved protocol with the University of Illinois Institutional 116 Animal Care and Use of Animals Committee (IUCAC #06081). 117 118 Fiber degradation analysis 119 Relative biomass (switchgrass) degradation during rumen-incubation was 120 determined by dry matter, organic matter, neutral detergent fiber (NDF), acid detergent 121 fiber (ADF), and acid detergent lignin (ADL) analysis. NDF, ADF and ADL were 122 determined using the procedures of Goering and Van Soest (Goering & Van Soest, 1970). 123 Cellulose content was estimated as ADF – ADL and hemicellulose as NDF – ADF. 124 125 DNA extraction and 16S rRNA gene amplification 126 Total microbial genomic DNA was extracted from 100 mg of the non-incubated 127 control sample and from each rumen-incubated biomass sample using a FastDNA SPIN 128 Kit for Soil (MP Biomedical, Solon, OH) according to the manufacturer’s instructions. 129 Extracted DNA was quantified with a spectrophotometer (Nanodrop ND1000; Thermo 130 Scientific, USA). The hypervariable V6 to V8 region of the 16S ribosomal RNA (rRNA) 131 gene was amplified from the environmental DNA using the primer set 926F/1392R (926F: 132 5’- cct atc ccc tgt gtg cct tgg cag tct cag AAA CTY AAA KGA ATT GRC GG- 3’ and 133 1392R: 5’ - cca tct cat ccc tgc gtg tct ccg act cag xxxxx ACG GGC GGT GTG TRC – 3’) 134 described by Engelbrektson et al. (Engelbrektson et al, 2010). Primer sequences were 135 modified by the addition of 454 A or B adapter sequences (lower case). In addition, the 136 reverse primer included a 5 bp barcode, indicated by xxxxx in the reverse primer 137 sequence above, for multiplexing of samples during sequencing. The barcode sequence 138 for each sample is listed in Table 1. The generated raw reads were deposited in NCBI’s 139 Short Read Archive under the accession number SRP042121. 140 141 Twenty-microliter PCR reactions were performed in duplicate and pooled to minimize 142 PCR bias. PCR reaction was performed using 0.4 µl Advantage GC 2 Polymerase Mix 143 (Advantage-2 GC PCR Kit, Clontech), 4 µl 5x GC PCR buffer, 2 µl 5 M GC Melt 144 Solution, 0.4 µl 10 mM dNTP mix (MBI Fermentas), 1.0 µl of each 25 nM primer, and 145 10 ng sample DNA. The thermal cycler protocol was 95°C for 3 min, 25 cycles of 95°C 146 for 30 s, 50°C for 45 s, and 68°C for 90 s, and a final 10 min extension at 68°C. PCR 147 amplicons were purified using Solid Phase Reversible Immobilization (SPRI) beads and 148 quantified using a Qubit flurometer (Invitrogen). Samples were diluted to 10 ng/µl and 149 mixed in equal concentrations. Emulsion PCR and sequencing of the PCR amplicons 150 were performed following the Roche 454 GS FLX Titanium technology instructions 151 provided by the manufacturer. 152 153 Pyrotag sequence analysis 154 Pyrosequencing data from the 20 samples were demultiplexed and processed 155 using QIIME version 1.7.0 (Caporaso et al, 2010) according to the standard operating 156 procedure described at http://qiime.org/tutorials/tutorial.html. Primers and barcodes were 157 removed before the raw reads were quality filtered. Sequences were removed if they had 158 long homopolymeric regions (> 6 nt), were smaller than 200 nt, had quality scores lower 159 than 25, or if they were identified as being chimeric. This resulted in a total of 198,037 160 high quality 16S rRNA gene sequences. Sequences generated from each of the biological 161 duplicates were combined prior to sequence analysis to account for biological variation. 162 163 OTU-based sequence analysis 164 Sequences were clustered into operational taxonomic units (OTUs) at a 97 % 165 sequence identity cut-off using UCLUST (Edgar, 2010). The most abundant sequence of 166 each OTU was picked as representative sequence. Singleton and doubleton abundance, 167 Shannon, Simpson, Simpson reciprocal, and Chao1 estimators were calculated using the 168 QIIME software. Representative sequences were aligned using the PyNAST algorithm 169 and the alignment was filtered to remove common gaps. Following the quality filtering 170 and grouping steps, 7,168 unique sequences (representing 198,037 total sequences) were 171 used for OTU analyses. QIIME scripts were used to calculate α-diversity metrics for 172 OTU representative sequences and to generate a Principal Coordinate Analysis plot 173 according to the standard operating procedure described at http://qiime.org/tutorials/ 174 tutorial.html. 175 176 Taxonomic classification of unique representative sequences 177 Taxonomic classification of the final set of representative sequences was 178 performed using QIIME. Each sequence was normalized to contain six taxonomic levels, 179 ranging from the domain to the genus level. 180 181 Scanning electron microscopy 182 For scanning electron microscopy (SEM) pieces of switchgrass taken from the 183 cow rumen as described above were immediately fixed with glutaraldehyde. The samples 184 were washed several times with phosphate buffered saline, treated with 1 % OsO4 for 1 h 185 on ice, prior to dehydration and critical point drying using a Tousimis Autosamdri-815 186 critical point dryer. The dried samples were then mounted on precut brass sample stubs 187 using double sided carbon tape and sputter coated with approximately 30 Angstrom of 188 Au/Pd. SEM imaging was performed on a Hitachi S-5000 microscope at 10 kV 189 accelerating voltage. 190 191 Results:

192 Temporal changes in α-diversity of fiber-adherent rumen microbiome 193 To determine α-diversity within the fiber-adherent community the quality-filtered 194 pyrotag reads of each sample were clustered into OTUs at a sequence identify level of 195 97 %. This resulted in a total of 7,168 distinct OTUs, with a range of 788 to 2,758 OTUs 196 observed for each community. Table 2 summarizes the number of raw reads, quality 197 filtered reads and OTUs that were obtained for each time point. Of the observed 7,168 198 OTUs, 7,005 (97.7 %) were determined to be unambiguously of bacterial origin, while 199 0.2 % (17 OTUs) were assigned to be archaeal (Table 3). Table 4 summarizes the number 200 of OTUs unique to each time point sampled. A complete list of all OTUs observed during 201 this study and an OTU count for each sample is provided in Table 5. Analysis of the 202 successional appearance of OTUs suggests an initial increase in community richness 203 within the first 30 min of rumen incubation (Tables 2 & 3) and a second increase in 204 richness of the bacterial community associated with the switchgrass samples subjected to 205 rumen-incubation for 16, 24, and 48 hours (Table 3). This finding is supported by the 206 calculated rarefaction curves (Figure 1) and Chao 1 indices (Table 6). Likewise the 207 Shannon and Simpson Reciprocal indices increased noticeably for all rumen-incubated 208 samples (Table 6). An average Good’s coverage estimator of 94.9 % was calculated for 209 all samples, with 92 % (6 h) being the lowest and 98.6 % (non-incubated switchgrass) 210 being the highest (Table 6). 211 212 Temporal changes in β-diversity of fiber-adherent rumen microbiome 213 Principal coordinates analysis comparing the temporal dynamics of switchgrass- 214 associated microbiota indicated that the community associated with the air-dried 215 switchgrass sample (0 h) was distinct from the microbiomes of all rumen-incubated 216 samples (Figure 2). After 30 min of rumen incubation the switchgrass-associated 217 microbiota exhibited clearly noticeable changes: for example, members of the Archaea, 218 which were essentially absent in the non-incubated microbiome, were detected in all 219 rumen-incubated samples. In total, 17 distinct archaeal OTUs were detected (Table 3), 220 with 14 (82 %) of them categorized as Methanobacteriaceae, a family belonging to the 221 methanogenic Euryarchaeota (Table 5). Of the reads derived from the Archaea, the 222 majority (≥ 76 %) was recruited for each sample (median 88 %) by two OTUs both 223 categorized as Methanobrevibacter following rumen incubation. Abundance of members 224 belonging to the genus Methylobacterium was notably high in the samples retrieved after 225 30 min of rumen incubation. The bacterial phylum Bacteroidetes, represented primarily 226 by members of the Sphingobacteria in the 0 h sample, increased up to five fold 227 throughout rumen incubation, mainly through large increases in Bacteroidia populations 228 and in particular of increases in members belonging to the genus Prevotella (Tables 7, 8 229 & 9). This overall increase in Bacteroidetes was accompanied by increased abundance of 230 Cyanobacteria, Fibrobacteres, Spirochaetes and Tenericutes and decreases in 231 Actinobacteria (i.e. Kineosporiaceae and Microbacteriaceae), Firmicutes and 232 Proteobacteria abundance (Table 9). 233 234 Observed decreases in Firmicutes were driven by a 460-fold reduction in the Bacilli 235 (mostly Bacillus spp.), which initially accounted for 97 % of the Firmicutes phylum and 236 63 % of all bacteria prior to rumen incubation. Decreases in Bacilli were partially offset 237 by large increases in the Clostridia including the Lachnospiraceae (particularly 238 Butyrivibrio spp., Pseudobutyrivibrio spp. and Clostridium spp.) and the 239 Ruminococcaceae (almost entirely Ruminococcus spp.) families. The Clostridia 240 continued to increase to become the predominant class and made Firmicutes the 241 predominant phylum after a brief peak in Proteobacteria abundance within the initial 30 242 min of rumen-incubation (Figure 3, Tables 7 & 9). The switchgrass-associated microbiota 243 stabilized at the phylum level after 1 h of rumen incubation (Figure 3), and only modest 244 phylum- and class-level changes were observed for most taxonomic groups between 1 245 and 72 h of rumen incubation (Tables 7 & 9). The only exceptions to this observation 246 were the Fibrobacteres and Spirochaetes, which both increased considerably in 247 abundance in samples collected between 6 and 16 h, peaking around 24 h before 248 returning to their 1 h levels after a total of 72 h of rumen-incubation (Figure 3, Table 9). 249 Each of these phylum-level changes was driven by a single genus: the Fibrobacteres 250 driven exclusively by Fibrobacter, while changes in the Spirochaetes were driven by 251 Treponema (Table 8). 252 253 Biomass-degradation coordinated with microbial colonization 254 Biomass degradation following rumen incubation appeared to be biphasic; each 255 phase corresponding to notable changes in microbial communities. Biomass degradation 256 began rapidly following rumen incubation with 13 % of the total biomass being degraded 257 within 30 min (Figure 4). Biomass-degradation then appeared to stall between 30 min and 258 4 h, with no significant change to biomass during this period. Corresponding to this 259 period, we observed noticeable increases in Methanobrevibacter spp. and changes in 260 some bacterial taxa. The Butyrivibrio spp. increased, while Prevotella spp. decreased 261 between 30 min and 4 h. Following 4 h, biomass degradation proceeded somewhat 262 linearly at a rate of ~0.81 % biomass h-1 for the next 20 h of incubation, before gradually 263 slowing over the following two measurements (~0.41 % biomass h-1 between 24 and 48 h 264 and 0.17 % biomass h-1 between 48 and 72 h). The increase in rate of biomass 265 degradation between 4 h and 24 h occurred simultaneously with increased numbers of 266 reads assigned to Fibrobacter and Treponema species (Figure 4, Table 8). Biomass- 267 degradation and colonization of switchgrass was also detectable by SEM with significant 268 fiber degradation being visible for samples incubated within the rumen for 24 h and 269 longer (Figure 5). 270 271 Discussion: 272 In the present study we observed temporal shifts in the plant biomass-associated 273 microbiota and in the rates of biomass degradation during the in situ rumen-incubation of 274 dried switchgrass. Changes in the microbiome were in particular prevailing immediately 275 within the first 30 min and after 4 h of rumen incubation. These observations are 276 consistent with analogous trials on rumen-incubated fresh perennial ryegrass, where 277 denaturing gradient gel electrophoresis (DGGE)-based analyses identified discrete 278 microbial profiles at 0-2 h and after 4 h onwards (Huws et al, 2013). Notable similarities 279 and differences were observed in several findings between these two studies. Consistent 280 with our findings, Huws and colleagues reported increases in richness and diversity 281 following rumen incubation (Huws et al, 2013). The authors of the former study also 282 found the major ruminal genus Prevotella (Stevenson & Weimer, 2007) increased after 4 283 h. In our study, Prevotella spp. growth was biphasic, increasing dramatically within the 284 first 30 min following rumen incubation and decreasing between 30 min and 4 h, before 285 increasing again. In our study, Prevotella spp. were most abundant at 6 h and did not 286 return to 4 h levels until after 24 h of incubation. In contrast to our observations, Huws 287 and colleagues reported a loss of a band corresponding to Treponema spp. after 4 h 288 rumen incubation (Huws et al, 2013), whereas we saw great increases in this genus 289 between 6 h and 16 h. This discrepancy likely reflects differences in analytical techniques 290 (DGGE vs. next-generation sequencing), where loss of a single Treponema band may not 291 reflect all members of that taxonomic group. Direct sequencing of 16S rRNA amplicons 292 also facilitated the detection of a significant increase of Methylobacterium in switchgrass 293 samples retrieved after 30 min of rumen incubation (Figure 4). Members of the genus 294 Methylobacterium are strictly aerobic, can utilize C1 compounds such as formate, 295 methanol and methaylamine and a variety of C2 (including acetate), C3, and C4 296 compounds (Green, 2006) as sole carbon source, and have been found in association with 297 a variety of plants (Corpe, 1985). We hypothesize that members of the Methylobacterium 298 were associated with the switchgrass fibers and introduced into the rumen, where they 299 metabolized common fermentation intermediates, such as formate and acetate (Sabine 300 and Johnson, 1964; Hungate et al., 1970), until all oxygen that might have been 301 introduced with the ground switchgrass was depleted. Overall many similarities exist 302 between the two studies and may indicate that the temporal changes in microbiota during 303 the first 30 min following rumen-incubation and then after 4 h may be common features 304 of rumen-microbial digestion among multiple plant species in various states (fresh vs. 305 dried). Changes in the microbiota corresponded with differences in the rates of 306 switchgrass degradation. The rate of switchgrass degradation was greatest within the first 307 30 min of rumen-incubation during which 13 % of the total biomass was lost. This may 308 reflect rapid utilization of soluble sugars and other easily fermentable nutrients that were 309 unabated by proximal H2 partial pressures. However, the rate of degradation stalled 310 between 30 min and 4 h. During this period we observed a dramatic increase in 311 methanogen density such that the total abundance of methanogens at 4 h post-incubation 312 increased 3.3-fold compared to 30 min incubation. We hypothesize that these two 313 observations were linked and related to a dramatic increase in proximal H2 partial 314 pressure caused by the rapid initial fermentation of easily accessible plant 315 polysaccharides. Methanogen numbers swelled, most likely in response to the 316 hypothesized high H2 partial pressures and were eventually able to overcome these and 317 create an environment that was favorable to the energetically-efficient growth of the 318 fibrolytic microbial consortia, as has been proposed based on in vitro evidence (Ishii et al, 319 2006; Stams, 1994). Although the community was largely established by 1 h, as 320 evidenced by less dramatic changes in the overall microbiota 16S rRNA profile following 321 this period, it was not until after 4 h before their collective metabolism was operating 322 efficiently. 323 324 In summary the findings presented here support the hypothesis that the hydrogenotrophic 325 metabolism of methanogens is an essential part of the fibrolytic activity of the rumen 326 ecosystem and is essential for the degradation of the more recalcitrant plant 327 polysaccharides by rumen microbes. Additional in situ work during which methane 328 emission, fiber composition and community are monitored more stringently will be 329 essential for obtaining a better holistic understanding of the rumen microbiome and the 330 contribution of particular community members to the fibrolytic and methanogenic rumen 331 phenotype. 332 333 Conflict of Interest Statement: 334 The authors declare that they have no conflict of interest. 335 336 Authors’ Contribution: 337 Conceived and designed the experiments: MH, ST, and RIM. Performed the experiments: 338 MH, HP, RIM. Generated and analyzed the data: MH, HP, ML, SM, AS, BK, MA, ST, 339 and CJY. Wrote the paper: MH, HP, ML, and CJY. 340 341 Acknowledgements: 342 The work conducted by the U.S. Department of Energy Joint Genome Institute is 343 supported by the Office of Science of the U.S. Department of Energy under Contract No. 344 DE-AC02-05CH11231. 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438 439 440 Figure Legends: 441 442 Figure 1. Rarefaction curves calculated from 16S rRNA pyrotag data of switchgrass- 443 adherent microbiomes after rumen-incubation. 0 h indicates the non-incubated 444 switchgrass sample (control). 445 446 Figure 2. Principal coordinate analysis or switchgrass-associated microbiomes. Each 447 point corresponds to one of nine rumen-incubated samples (0.5, 1, 2, 4, 6, 16, 24, 48, and 448 72 h) and one control sample (0 h). The percentage of variation explained by the plotted 449 principal coordinates is indicated on the axes. 450 451 Figure 3. Effect of rumen incubation on relative abundance of bacterial and archaeal 452 phyla of switchgrass-adherent microbiome. Relative OTU abundance of rarefied 16S 453 rRNA gene 454-pyrotag data. Taxonomy was assigned based on Greengenes (McDonald 454 et al, 2012). 455 456 Figure 4. Microbial succession and biomass-degradation during rumen-incubation. Heat 457 map show succession of genera recruiting >0.5 % of the generated sequences. Line 458 graphs show the relative dry mater change during rumen-incubation. (A) Upper panel 459 shows genera with >10 % relative abundance, lower panel shows genera with a relative 460 abundance between 2-10 %; (B) Upper panel shows genera with relative abundance 461 between 1-2 %, lower panel shows genera with a relative abundance between 0.5-1 %. 462 463 Figure 5. Scanning Electron Microscopy of air-dried and rumen-incubated switchgrass 464 samples and adherent microorganisms. Scale bars indicate 0.5 µm. 465 Figure 1.TIF Figure 2.TIF Figure 3.TIF Figure 4.TIF Figure 5.JPEG Copyright of Frontiers in Microbiology is the property of Frontiers Media S.A. and its content may not be copied or emailed to multiple sites or posted to a listserv without the copyright holder's express written permission. However, users may print, download, or email articles for individual use.