Transcriptomic characterization of Caecomyces churrovis: a novel, non-rhizoid-forming lignocellulolytic anaerobic fungus
© The Author(s) 2017
Received: 3 August 2017
Accepted: 12 December 2017
Published: 20 December 2017
Anaerobic gut fungi are the primary colonizers of plant material in the rumen microbiome, but are poorly studied due to a lack of characterized isolates. While most genera of gut fungi form extensive rhizoidal networks, which likely participate in mechanical disruption of plant cell walls, fungi within the Caecomyces genus do not possess these rhizoids. Here, we describe a novel fungal isolate, Caecomyces churrovis, which forms spherical sporangia with a limited rhizoidal network yet secretes a diverse set of carbohydrate active enzymes (CAZymes) for plant cell wall hydrolysis. Despite lacking an extensive rhizoidal system, C. churrovis is capable of growth on fibrous substrates like switchgrass, reed canary grass, and corn stover, although faster growth is observed on soluble sugars. Gut fungi have been shown to use enzyme complexes (fungal cellulosomes) in which CAZymes bind to non-catalytic scaffoldins to improve biomass degradation efficiency. However, transcriptomic analysis and enzyme activity assays reveal that C. churrovis relies more on free enzymes compared to other gut fungal isolates. Only 15% of CAZyme transcripts contain non-catalytic dockerin domains in C. churrovis, compared to 30% in rhizoid-forming fungi. Furthermore, C. churrovis is enriched in GH43 enzymes that provide complementary hemicellulose degrading activities, suggesting that a wider variety of these activities are required to degrade plant biomass in the absence of an extensive fungal rhizoid network. Overall, molecular characterization of a non-rhizoid-forming anaerobic fungus fills a gap in understanding the roles of CAZyme abundance and associated degradation mechanisms during lignocellulose breakdown within the rumen microbiome.
Anaerobic gut fungi are robust degraders of plant biomass in the guts of ruminants and other large monogastric herbivorous mammals . They have also been identified using microscopy and molecular methodologies in the digestive tract of herbivorous reptiles . Due to the large amount of biomass-degrading enzymes that these organisms secrete [3–5], they have potential for application in the production of lignocellulose-derived chemical products . Most known genera (Anaeromyces, Buwchfawromyces, Neocallimastix, Oontomyces, Orpinomyces, Pecoramyces, Piromyces) of gut fungi have an extensive network of penetrating rhizoids (tapering mycelia) that aid, alongside enzymatic activity, in biomass colonization and deconstruction [7–11]. However, two known genera within the clade of anaerobic fungi (Caecomyces, Cyllamyces) do not produce rhizoidal networks, but form a limited system simply capable of attaching to plant biomass [7, 12]. However, like their rhizoidal counterparts, non-rhizoid producing gut fungi are proficient degraders of crude plant biomass. This raises the possibility that there are differences between the diversity of enzymes employed by rhizoid-forming vs. non-rhizoid-forming fungi, and/or the mechanisms they use for enzymatic degradation of lignocellulose.
While there is a general lack of genomic information for the Neocallimastigomycota, recently five complete genomes and transcriptomes have been published for rhizoid-forming anaerobic fungi—two representatives from the Piromyces genus, one each from Anaeromyces and Neocallimastix , and one from Orpinomyces  (recently reclassified as Pecoramyces ). Interestingly, this wealth of sequencing data has revealed that anaerobic fungi can draw from two modes of biomass-degradation via the secretion of freely diffusive enzymes as well as via fungal cellulosomes (complexes of enzymes tethered together for synergistic action) . However, at the present time, no high-resolution transcriptomic or genomic information has been reported for non-rhizoid-forming isolates. This precludes insight into the enzymatic machinery of non-rhizoid-forming fungi, or the mode of biomass degradation that they favor.
Here, we describe a novel species of non-rhizoid-forming fungi belonging to the Caecomyces genus isolated from the fecal pellets of a Navajo Churro sheep collected in 2015. While other Caecomyces isolates have been described using morphological and phylogenetic analyses [15, 16], including some analysis of the cellulolytic enzyme activity , extensive genomic or transcriptomic sequencing has not been completed. By assembling the first sequenced transcriptome for an anaerobic gut fungus within the Caecomyces genus, C. churrovis, our analysis enabled us to identify the range of CAZymes available to a non-rhizoidal genus and test the null hypothesis that additional degradation mechanisms, mechanical or enzymatic, are not required for dissolution of plant biomass. This isolated fungal strain was assessed for its ability to grow on a range of substrates, and demonstrated a greater preference for soluble substrates compared to other rhizoid-forming strains that have been analyzed to date . Transcriptome assembly and analysis identified a broad range of CAZymes within the genome, including a relative abundance of carbohydrate esterase and hemicellulase (GH 11/12, 43) transcripts. Comparison to other sequenced gut fungal isolates also revealed a greater reliance on free enzymes rather than enzymes bound in fungal cellulosome complexes.
Results and discussion
Isolation and molecular classification of C. churrovis
C. churrovis is capable of growth on crude biomass and fermentable sugars
Among soluble sugars, the highest overall pressure production was observed during growth on cellobiose (15.37 ± 0.42 psig) and glucose (15.0 ± 0.20 psig), with lower pressure observed during growth on fructose (8.93 ± 1.1 psig) suggesting that greater growth and metabolic activity occurs on glucose and cellobiose (Fig. 3). This may be related to regulation mechanisms behind the production of biomass-degrading enzymes, as glucose has been shown to function as a carbon catabolite repressor for CAZymes , resulting in fewer cellular resources being diverted to produce enzymes that are not necessary for growth on these substrates. Fungal growth was not observed on the five-carbon sugars xylose and arabinose, or six-carbon sugars galactose and mannose, despite the presence of these sugars in plant cell walls. Cellobiose—a breakout product of cellulose—was the only disaccharide that supported growth, as no growth was observed on maltose and sucrose.
For polymeric substrates, minimal growth was detected on purified crystalline cellulose (Avicel® and Sigmacell) with total accumulated pressure only 1.2–3.5 psig greater than total accumulated pressure of blank cultures in the absence of a carbon source. However, the lack of growth on crystalline cellulose may be linked to the lack of rhizoids produced by C. churrovis. The small particle size (50 µm) of these crystalline cellulose substrates compared to milled plant biomass (4 mm) may lead to more dense substrate packing, preventing access of the gut fungus to the cellulose beyond the top surface layer. In contrast, rhizoidal networks produced by other gut fungi likely penetrate the cellulose, disrupting the packing and improving access of zoospores and secreted enzymes to exposed cellulose chains. C. churrovis growth was observed on xylan for some cultures, but was inconsistent and resulted in large error in growth rate measurements (Fig. 3). Growth rates observed on plant biomass substrates varied significantly with net-specific growth rates on corn stover (0.039 ± 0.0002 h−1) significantly higher (P = 1.4 × 10−4) than on switchgrass and reed canary grass (0.030 ± 0.0005 h−1 and 0.028 ± 0.003 h−1, respectively), and no growth observed on alfalfa stems. This observation is consistent with the cell wall composition of alfalfa stems, which have a greater relative pectin content compared to the other grasses , which may hinder fungal growth. The greatest pressure was observed during growth on corn stover (12.4 ± 0.10 psig), while lower total pressure was measured on reed canary grass and switchgrass (8.27 ± 0.76 and 9.0 ± 0.35 psig, respectively). This suggests that differing composition of the plant material in terms of lignin content and sugar composition [25, 26] may impact growth of C. churrovis. Corn stover comprises 32–36% glucan  while reed canary grass and switch grass comprise 20.9–26.5 and 27.3–32.2% cellulose, respectively . This greater glucan concentration in corn stover may have resulted in greater maximum pressure accumulation during growth on this substrate.
Comparative transcriptomic analysis of C. churrovis against rhizoid-forming gut fungi
C. churrovis transcriptome sequencing and annotation summary
Transcriptome size (bp)
Number of transcripts
Average length (bp)
Number of predicted genes (transcripts less isoforms)
Number of clusters
Number of reads
% with EC number
% with BLAST hits
% with gene ontology
% with InterPro scan
Transcriptomic Comparison of C. churrovis to other Anaerobic Gut Fungi
# C. churrovis transcripts aligned
% C. churrovis transcriptome
# transcripts matched
% transcriptome aligned to
Comparison of cellulose machinery across four gut fungal strains
Number of transcripts (# dockerin containing transcripts)a
Comparison of CAZyme machinery reveals a dependency on free enzymes in C. churrovis
Interestingly, C. churrovis harbors fewer transcripts for polysaccharide deacetylases (8.2% of CAZymes) compared to the other rhizoid-forming fungi where it is the most abundant CAZyme family (14.3–19.2%). In contrast, the highest abundance accessory enzymes in C. churrovis were carbohydrate esterases (9.2% of CAZymes) containing SGNH hydrolase domains. A similar proportion of carbohydrate esterases was observed in A. robustus, but a smaller proportion was observed in N. californiae and P. finnis. These enzymes aid in the digestion of plant wall polysaccharides by removing acetylation and improving hydrolysis . A greater proportion of the pectin lyase transcripts were also identified in C. churrovis (8.8% compared to 1.7–5.4%). The identification of many putative pectin degrading enzymes suggests that C. churrovis should be able to grow on pectin-rich grasses like alfalfa. However, this activity was not observed in C. churrovis, despite reports that rhizoid-forming fungi can subsist on alfalfa stems . This growth discrepancy may be due to the fact that alfalfa stems formed a gelatinous layer on top of the plant material after autoclaving, forming an effective “barrier” against penetration by non-rhizoid-forming fungi like Caecomyces. It is therefore possible that this limited the ability of the fungus to access and colonize the plant material.
As a mechanism to more efficiently degrade plant biomass, anaerobic gut fungi have been described to form complexes of CAZymes (fungal cellulosomes, Additional file 1: Figure S3), bringing the activities of multiple enzymes in closer proximity to each other. Recently, structural scaffoldin proteins have been discovered that mediate this complex formation by binding to non-catalytic fungal dockerin domains that are distinct from previously described bacterial dockerins . Given the similarity across gut fungal CAZymes, we hypothesized that some (or all) of this machinery might be shared between previously sequenced fungi and C. churrovis. Therefore, transcripts encoding for scaffoldin proteins in C. churrovis were identified by aligning the amino acid sequence of transcripts identified as scaffoldins in P. finnis  to the C. churrovis transcriptome using tblastn alignment. This revealed 38 transcripts with an alignment E-value of 0 (Additional file 1: Table S2), indicating that the scaffoldin machinery that forms fungal cellulosome complexes is actively transcribed under the substrate conditions encompassed in the transcriptome of C. churrovis. However, extensive Hidden Markov Model (HMM) analysis was not completed as done in previous work , largely due to the difficulties of applying such techniques in the absence of corresponding high-resolution genomes.
We also identified and compared the CAZyme transcripts containing non-catalytic fungal dockerin domains (NCDDs) (also referred to as CBM10s) to identify the protein components of the fungal cellulosome in C. churrovis (Table 3). While the general diversity of NCDD containing CAZyme transcripts was relatively consistent across strains with 45–55% cellulases, 36–47% hemicellulases, and 6–10% accessory enzymes, the percent of all CAZymes in C. churrovis with NCDDs was significantly lower compared to other strains. In C. churrovis, of the 512 CAZyme transcripts identified 77 also contained a fungal dockerin domain, representing 15% of all CAZyme transcripts. By comparison, the fraction of cellulosome-associated CAZymes in C. churrovis is much lower compared to rhizoid-forming A. robustus, N. californiae, and P. finnis, in which the dockerin-containing CAZyme transcripts represent 27.9, 29.3, and 31.4% of all CAZyme transcripts, respectively [5, 13]. This suggests that C. churrovis places greater emphasis on secreted un-complexed, free enzymes to attack plant biomass and release fermentable sugars compared to rhizoid-forming anaerobic fungi. C. churrovis does not maintain as much physical contact to the surface of plant material compared to rhizoid-forming strains that penetrate and intertwine with it. This dependence on free enzymes may maximize the area of the plant surface acted on by secreted CAZymes, especially if scaffoldin proteins are anchored to the fungal cell membrane or cell wall.
To test this finding, the activity of C. churrovis CAZymes on carboxymethyl cellulose (CMC), xylan, and pectin from cellulosome complexes was compared to the activity of all enzymes present in the fungal culture supernatants. Fungal cellulosomes were isolated through cellulose precipitation as previously described [5, 36]. This method enriches for cellulosome complexes rather than free enzymes. In contrast, the activity of the fungal supernatant contains cellulosome and free enzyme activities, in addition to non-cellulolytic proteins. Here, the enzyme-rich culture supernatant from C. churrovis possessed the highest activity relative to the prepared cellulosome compared to the other fungi tested (Fig. 4). Conversely, the cellulosome preparations of P. finnis, N. californiae, and A. robustus exhibited much greater activity than their corresponding culture supernatants containing a mixture of complexed and free enzymes. This finding is consistent with the hypothesis that C. churrovis is more dependent on free enzyme activity for the breakdown of cellulosic substrates compared to other rhizoid-forming genera of gut fungi that transcribe a higher fraction of cellulosome-associated enzymes. Furthermore, the activity on CMC, xylan, and pectin demonstrates the wide range of cellulolytic and hemicellulolytic enzyme activities as determined from transcriptomics.
Among anaerobic gut fungi, the Caecomyces genus represents an interesting opportunity to identify the role of gut fungal enzymes in their native microbiome in the absence of extensive, invasive rhizoidal growth. Here, we have characterized the growth of the novel isolate, C. churrovis across plant biomass substrates ranging in complexity and composition. C. churrovis demonstrated the most rapid growth on free sugars like glucose, cellobiose, and fructose and no growth on other sugars that are derived from plant biopolymers. Despite the lack of invasive rhizoids, C. churrovis was capable of growth on complex plant biomasses reed canary grass, corn stover, and switchgrass. Sequencing and assembly of the first transcriptome for an anaerobic gut fungus within the Caecomyces genus identified a broad array of CAZymes, including an increased diversity of hemicellulases compared to its rhizoid-forming counterparts. Without the mechanical disruption provided by rhizoidal growth , the suite of enzymes secreted by C. churrovis was still sufficient for hydrolysis of crude plant material. Cellulosome complex forming scaffoldin proteins were identified in the transcriptome, but a smaller proportion of CAZyme transcripts containing NCDDs suggest a greater dependence on free enzymes for plant biomass degradation compared to rhizoid-forming gut fungal genera. Enzyme activity assays supported this hypotheses as C. churrovis cellulosome preparations showed the least improved biomass-degrading activity relative to fungal culture supernatants. Here, our study of a non-rhizoid-forming gut fungus highlights the capabilities of gut fungal enzymes as a mechanism for lignocellulose hydrolysis and, in the case of C. churrovis, a greater reliance on free enzymes.
Isolation and culture maintenance
Strictly anaerobic, aseptic techniques and an incubation temperature of 39 °C were used throughout for fungal isolation and culture maintenance. The headspace gas was 100% CO2 and the antibiotic, chloramphenicol, at a final medium concentration of 100 µg mL−1, was used in all liquid culture media, but not in agar containing roll tubes. C. churrovis (IF 553979) was isolated from fresh fecal pellets from the Navajo Churro sheep enclosure at the Santa Barbara Zoo (Santa Barbara, CA). Fresh fecal material was returned to the laboratory within 2 h of collection, ground, and suspended into culture Medium C . Resuspended fecal material was diluted in a 10-fold series and 1 mL aliquots of the higher dilutions were used to inoculate anaerobic Hungate tubes containing Medium C and sterilized, 4-mm-milled reed canary grass. Cultures that demonstrated fungal growth and the absence of bacterial contamination were sustained on reed canary grass through routine anaerobic transfers into antibiotic containing culture media. Axenic cultures were obtained using roll tubes (25-mL serum tubes coated with 5 mL of solidified Medium C containing 2% agar and 0.5% cellobiose as the sole carbon source) inoculated with 0.1 mL of actively growing culture. Inoculated roll tubes were incubated for 2–3 days, after which isolated single colonies were selected by cutting them out of the agar and transferring to grow on reed canary grass in fresh, anaerobic liquid culture tubes. This procedure was performed in a Styrofoam box under a constant flow of CO2 to maintain anaerobic conditions. This process of colony selection, picking, and culture was completed three times for each strain of gut fungus to ensure selection of a single, isolated strain. These axenic cultures were routinely grown in 10-mL batch cultures of Medium C containing ground reed canary grass (4-mm particle size) in 15-mL anaerobic Hungate tubes. The antibacterial antibiotic was withdrawn from culture media after the single colony isolation process, once it was absolutely certain that cultures did not contain contaminating bacteria. Cultures were routinely transferred to new media every 3–5 days to continue growth. Cultures were also stored cryogenically, as described by Solomon, Henske et al. .
Phylogenetic analysis of isolated fungi
Phylogenetic analysis was completed by sequencing the internal transcribed spacer (ITS) region for each of the isolated fungi. ITS sequences were PCR amplified using the previously described JB206 and JB205 primers . The amplified DNA was sequenced and the ITS1 region was primarily employed in phylogenetic analysis. ITS1 or full ITS sequences were obtained for other anaerobic gut fungi across all known genera from sequences deposited in GenBank [29, 39]. The phylogenetic tree was created using Molecular Evolutionary Genetic Analysis (MEGA) software version 6.0 . Sequences were aligned using the Clustal Omega multiple sequence alignment method [41, 42], and the alignment was used to construct phylogeny using the maximum parsimony method. To test the confidence of the phylogeny, a bootstrap method was used with 1000 replications. Trees were edited for display using the Interactive Tree of Life .
Helium ion microscopy
Helium Ion Microscopy was performed as described in Henske et al. . Briefly fungal cultures were chemically fixed with 2% glutaraldehyde (Sigma Aldrich) and dehydrated through step-gradients from 0 to 70% ethanol. The biomass was then washed with 100% ethanol and dried using critical point drying with an Autosamdri-815 (Tousimis, Rockville, MD) and CO2 as a transitional fluid. Dried samples were sputter-coated with conductive carbon and secondary electron images were obtained with an Orion helium ion microscope (Carl Zeiss Microscopy, Peabody, MA).
Growth analysis of C. churrovis
Growth curves of axenic fungal cultures were generated from Medium C grown cultures by measuring the pressure of fermentation gases during growth, which is a precise, indirect measure of fungal proliferation . Soluble substrates, glucose, fructose, galactose, xylose, arabinose, maltose, cellobiose, and sucrose were dissolved in water and sterile filtered. They were added to autoclaved Medium C to a final concentration of 5 g L−1. Carboxymethyl cellulose, Avicel®, Sigmacell (Sigma Aldrich), xylan (from corn stover, TCI Chemicals, Portland, OR), reed canary grass, corn stover, switchgrass, and alfalfa stems (USDA-ARS Research Center, Madison, WI) were added to a concentration of 10 g L−1 prior to autoclaving media. Pressure measurements were taken five times daily for 10 days. Effective net-specific growth rates were determined from the slope of pressure accumulation data plotted against fermentation time of 3 × replicate cultures during the phase of exponential gas accumulation.
RNA isolation for transcriptome acquisition
RNA was isolated from growing fungal cultures during the exponential growth phase using the Qiagen RNeasy Mini Kit (Qiagen, Valencia, CA), as previously described using the plant and fungi protocol with liquid nitrogen grinding and on-column DNase Digest . RNA was isolated from cultures grown on glucose, fructose, cellobiose, cellulose, and reed canary grass. The RNA quality was determined through measurement on an Agilent Tapestation 2200 (Agilent, Santa Clara, CA) to obtain RINe scores and the minimum RINe score for samples used in transcriptome acquisition was 8.9. The total RNA quantity was determined by using Qubit Fluorometric Quantitation (Qubit, New York, NY) using the high sensitivity RNA reagents.
RNA sequencing and transcriptome assembly
RNA was pooled prior to generation of the sequencing library using equal quantities of total RNA from each growth condition. After pooling libraries were created using an Illumina Truseq Stranded mRNA library prep kit (Illumina Inc., San Diego, CA) following the kit protocol. The transcriptome was sequenced using the UCSB Biological Nanostructures Laboratory core sequencing facility’s Illumina NextSeq. Coverage greater than 500 × was achieved and assembled and the transcriptome was assembled de novo using Trinity .
Transcriptome annotation and analysis
The transcriptome of C. churrovis was annotated as previously described using a combination of NCBI Blast, InterPro, Gene Ontology, and ortholog annotations . Blast annotation was completed using the NCBI standalone blast application to perform blastx against the NCBI non-redundant database downloaded on 11/25/2015  with an E-value cutoff of 10−3. Transcripts were then analyzed for protein domains using the BLAST2GO package  for alignment to sequences in the EMBL-EBI InterPro database before gene ontology  terms and enzyme commission  numbers were assigned. Antisense RNA (asRNA) sequences were identified based on the strand specificity of the library and orientation of alignments to BLAST hits. All transcripts were examined for orthology by comparing all possible open reading frames to the OrthoMCL database .
Gut fungal transcriptome sequence comparisons
Alignment of full gut fungal transcriptomes was completed using the standalone BLAST tool kit . BLAST databases were created from full transcriptome fasta files using the “makeblastdb” function. The blastn function was then used to align transcriptome fasta files to transcriptome databases. For identification of scaffoldin transcripts, amino acid sequences of four scaffoldin transcripts from Piromyces finnis  were aligned to the C. churrovis transcriptome nucleotide database using tblastn which aligns the amino acid sequences to the translated (in all frames) sequences in the database.
Cellulase activity assays
Fungal enzymatic activity on Carboxymethyl Cellulose (CMC) (Sigma Aldrich), xylan (from corn stover, TCI Chemicals, Portland, OR), and pectin (from citrus fruits, MP biomedicals) was measured essentially as described previously . Briefly, 50 µL of a 2% substrate solution in citrate–phosphate buffer (pH 6.5) was combined with 30 µL of the cellulosome fraction or supernatant. The reducing sugar concentration was measured by adding 60 µL of DNS to 30 µL of reaction and then heating the solution at 95 °C for 5 min. 36 µL of the completed DNS reaction were transferred to 160 µL of water and the absorbance was measured at 540 nm. Rates were calculated by comparing to a standard curve constructed from glucose, and by subtracting a blank measurement where blank buffer was added to the substrate. In all cases, samples were performed in triplicate, and all values were normalized by total protein as measured by a BCA protein assay kit (Pierce Biotechnology, Rockford, IL). Cellulosome fractions were prepared as previously described using cellulose to precipitate cellulosome complexes from fungal supernatant after growth for 6 days on reed canary grass in Medium C [5, 36].
JKH performed all experiments and analysis except for enzyme activity and microscopy. SPG and DK performed enzyme activity assay analysis. FJC performed light microscopy. CRS, VS, and JEE performed helium ion microscopy. JKH, JAS, MKT isolated the fungal strain. JKH and MAO conceived the project and wrote the manuscript. All authors read and approved the final manuscript.
Funding support for this work was provided by the Office of Science (BER), US Department of Energy (DE-SC0010352), the Institute for Collaborative Biotechnologies through grant W911NF-09-0001 from the US Army Research Office, and the National Science Foundation (MCB-1553721). A portion of this research used resources at the Environmental Molecular Sciences Laboratory a DOE Office of Science User Facility. The facility is sponsored by the Office of Biological and Environmental Research and operated under Contract No. DE-AC05-76RL01830. RNA sequencing data were generated in the UCSB and UCOP-operated Biological Nanostructures Laboratory within the California NanoSystems Institute (CNSI) and analyzed using the computational resources of the Center for Scientific Computing from the CNSI, MRL: an NSF MRSEC (DMR-1121053) and NSF CNS-0960316. The authors also thank Paul Weimer from the United States Department of Agriculture (USDA) for providing freshly milled biomass substrates. Prof Theodorou gratefully acknowledges funded contributions from Harper Adams University and UCSB which contributed to his sabbatical leave at UCSB. Evans, Smallwood, and Shutthanandan were supported by DOE-BER Mesoscale to Molecules Bioimaging Project #66382. Henske also acknowledges the Mellichamp Academic Initiative in Sustainability for fellowship support, and Gilmore was supported by the National Science Foundation Graduate Research Fellowship Program under Grant No. DGE 114085.
The authors declare that they have no competing interests.
US Department of Energy (DE-SC0010352). US Army Research Office, Institute for Collaborative Biotechnologies through grant W911NF-09-0001. National Science Foundation (MCB-1553721).
Illumina read data (.fastq files) used for the assembly of the C. churrovis transcriptome have been submitted to the NCBI Sequence Read Archive (SRA) under BioProject Number PRJNA393353 and BioSample Submission Number SUB2845331. The Internal Transcribed Spacer (ITS) region of C. churrovis used for phylogenetic analysis can be found on GenBank under Reference Number MF460993. C. churrovis was registered with Index Fungorum with the IF Number 553979.
Ethics approval and consent to participate
Springer Nature remains neutral with regard to jurisdictional claims in published maps and institutional affiliations.
Open AccessThis article is distributed under the terms of the Creative Commons Attribution 4.0 International License (http://creativecommons.org/licenses/by/4.0/), which permits unrestricted use, distribution, and reproduction in any medium, provided you give appropriate credit to the original author(s) and the source, provide a link to the Creative Commons license, and indicate if changes were made. The Creative Commons Public Domain Dedication waiver (http://creativecommons.org/publicdomain/zero/1.0/) applies to the data made available in this article, unless otherwise stated.
- Theodorou MK, Mennim G, Davies DR, Zhu WY, Trinci AP, Brookman JL. Anaerobic fungi in the digestive tract of mammalian herbivores and their potential for exploitation. Proc Nutr Soc. 1996;55:913–26.View ArticleGoogle Scholar
- Liggenstoffer AS, Youssef NH, Couger MB, Elshahed MS. Phylogenetic diversity and community structure of anaerobic gut fungi (phylum Neocallimastigomycota) in ruminant and non-ruminant herbivores. ISME J. 2010;4(10):1225–35.View ArticleGoogle Scholar
- Haitjema CH, Solomon KV, Henske JK, Theodorou MK, O’Malley MA. Anaerobic gut fungi: advances in isolation, culture, and cellulolytic enzyme discovery for biofuel production. Biotechnol Bioeng. 2014;111(8):1471–82.View ArticleGoogle Scholar
- Ljungdahl LG. The cellulase/hemicellulase system of the anaerobic fungus Orpinomyces PC-2 and aspects of its applied use. Ann N Y Acad Sci. 2008;1125:308–21.View ArticleGoogle Scholar
- Solomon KV, Haitjema CH, Henske JK, Gilmore SP, Borges-Rivera D, Lipzen A. Early-branching gut fungi possess a large, comprehensive array of biomass-degrading enzymes. Science. 2016;351:1192–5.View ArticleGoogle Scholar
- Gilmore SP, Henske JK, O’Malley MA. Driving biomass breakdown through engineered cellulosomes. Bioengineered. 2015;6(4):204–8.View ArticleGoogle Scholar
- Mountfort DO, Orpin CG. Anaerobic fungi: biology, ecology, and function. Boca Raton: CRC Press; 1994.Google Scholar
- Dagar SS, Kumar S, Griffith GW, Edwards JE, Callaghan TM, Singh R, Nagpal AK, Puniya AK. A new anaerobic fungus (Oontomyces anksri gen. nov., sp. nov.) from the digestive tract of the Indian camel (Camelus dromedarius). Fungal Biol. 2015;119(8):731–7.View ArticleGoogle Scholar
- Griffith GW, Callaghan TM, Podmirseg SM, Hohlweck D, Edwards JE, Puniya AK, Dagar SS. Buwchfawromyces eastonii gen. nov., sp. nov.: a new anaerobic fungus (Neocallimastigomycota) isolated from buffalo faeces. MycoKeys. 2015;9:11–28.View ArticleGoogle Scholar
- Hanafy RA, Elshahed MS, Liggenstoffer AS, Griffith GW, Youssef NH. Pecoramyces ruminantium, gen. nov., sp. nov., an anaerobic gut fungus from the feces of cattle and sheep. Mycologia. 2017;109(2):231–43.View ArticleGoogle Scholar
- Ho YW, Abdullah N, Jalaludin S. Penetrating structures of anaerobic rumen fungi in cattle and swamp buffalo. Microbiology. 1988;134(1):177–81.View ArticleGoogle Scholar
- Ozkose E, Thomas BJ, Davies DR, Griffith GW, Theodorou MK. Cyllamyces aberensis gen. nov. sp. nov., a new anaerobic gut fungus with branched sporangiophores isolated from cattle. Can J Bot. 2001;79(6):666–73.Google Scholar
- Haitjema CH, Gilmore SP, Henske JK, Solomon KV, Groot RD, Kuo A, Mondo SJ, Salamov AA, Labutti K, Zhao Z, et al. A parts list for fungal cellulosomes revealed by comparative genomics. Nat Microbiol. 2017;2:17087.View ArticleGoogle Scholar
- Youssef NH, Couger MB, Struchtemeyer CG, Liggenstoffer AS, Prade RA, Najar FZ. The genome of the anaerobic fungus Orpinomyces sp. strain C1A reveals the unique evolutionary history of a remarkable plant biomass degrader. Appl Environ Microbiol. 2013;79:4620–34.View ArticleGoogle Scholar
- Chen Y-C, Tsai S-D, Cheng H-L, Chien C-Y, Hu C-Y, Cheng T-Y. Caecomyces sympodialis sp. nov., a new rumen fungus isolated from Bos indicus. Mycologia. 2017;99(1):125–30.View ArticleGoogle Scholar
- Wubah DA, Fuller MS, Akin DE. Studies on Caecomyces communis: morphology and development. Mycologia. 1991;83(3):30–3310.View ArticleGoogle Scholar
- Hodrova B, Kopecny J, Kas J. Cellulolytic enzymes of rumen anaerobic fungi Orpinomyces joyonii and Caecomyces communis. Res Microbiol. 1998;149(6):417–27.View ArticleGoogle Scholar
- Schoch CL, Seifert KA, Huhndorf S, Robert V, Spouge JL, Levesque CA, Chen W, Fungal Barcoding C. Fungal Barcoding Consortium Author L: nuclear ribosomal internal transcribed spacer (ITS) region as a universal DNA barcode marker for fungi. Proc Natl Acad Sci USA. 2012;109(16):6241–6.View ArticleGoogle Scholar
- Brookman JL, Mennim G, Trincia P, Theodorou MK, Tuckwell DS. Identification and characterization of anaerobic gut fungi using molecular methodologies based on ribosomal ITS1 and 18S rRNA. Microbiology. 2000;146:393–403.View ArticleGoogle Scholar
- Tuckwell DS, Nicholson MJ, McSweeney CS, Theodorou MK, Brookman JL. The rapid assignment of ruminal fungi to presumptive genera using ITS1 and ITS2 RNA secondary structures to produce group-specific fingerprints. Microbiology. 2005;151(Pt 5):1557–67.View ArticleGoogle Scholar
- Nicholson MJ, McSweeney CS, Mackie RI, Brookman JL, Theodorou MK. Diversity of anaerobic gut fungal populations analysed using ribosomal ITS1 sequences in faeces of wild and domesticated herbivores. Anaerobe. 2010;16(2):66–73.View ArticleGoogle Scholar
- Li GJ, Hyde KD, Zhao RL, Hongsanan S, Abdel-Aziz FA, Abdel-Wahab MA. Fungal diversity notes 253–366: taxonomic and phylogenetic contributions to fungal taxa. Fungal Divers. 2016;78:1–237.View ArticleGoogle Scholar
- Theodorou MK, Davies DR, Nielsen BB, Lawrence MIG, Trinci APJ. Determination of growth of anaerobic fungi on soluble and cellulosic substrates using a pressure transducer. Microbiology. 1995;141(3):671–8.View ArticleGoogle Scholar
- Theodorou MK, Williams B, Dhanoa MS, McAllan AB, France J. A simple gas production method using a pressure transducer to determine the fermentation kinetics of ruminant feeds. Anim Feed Sci Technol. 1994;48(3–4):185–97.View ArticleGoogle Scholar
- Dien B, Jung H, Vogel K, Casler M, Lamb J, Iten L, Mitchell R, Sarath G. Chemical composition and response to dilute-acid pretreatment and enzymatic saccharification of alfalfa, reed canarygrass, and switchgrass. Biomass Bioenerg. 2006;30(10):880–91.View ArticleGoogle Scholar
- Pordesimo LO, Hames BR, Sokhansanj S, Edens WC. Variation in corn stover composition and energy content with crop maturity. Biomass Bioenerg. 2005;28(4):366–74.View ArticleGoogle Scholar
- Grabherr MG, Haas BJ, Yassour M, Levin JZ, Thompson DA, Amit I, Adiconis X, Fan L, Raychowdhury R, Zeng Q, et al. Full-length transcriptome assembly from RNA-Seq data without a reference genome. Nat Biotechnol. 2011;29(7):644–52.View ArticleGoogle Scholar
- Martin J, Bruno VM, Fang Z, Meng X, Blow M, Zhang T, Sherlock G, Snyder M, Wang Z. Rnnotator: an automated de novo transcriptome assembly pipeline from stranded RNA-Seq reads. BMC Genom. 2010;11(1):663–70.View ArticleGoogle Scholar
- NCBI Resource Coordinators. Database resources of the national center for biotechnology information. Nucleic Acids Res. 2016;44(D1):D7–19.View ArticleGoogle Scholar
- Finn RD, Attwood TK, Babbitt PC, Bateman A, Bork P, Bridge AJ, Chang HY, Dosztanyi Z, El-Gebali S, Fraser M, et al. InterPro in 2017-beyond protein family and domain annotations. Nucleic Acids Res. 2017;45(D1):D190–9.View ArticleGoogle Scholar
- Nicholson MJ, Theodorou MK, Brookman JL. Molecular analysis of the anaerobic rumen fungus Orpinomyces—insights into an AT-rich genome. Microbiology. 2005;151(Pt 1):121–33.View ArticleGoogle Scholar
- Brownlee AG. Remarkably AT-rich genomic DNA from the anaerobic fungus Neocallimastix. Nucleic Acids Res. 1989;17(4):1327–35.View ArticleGoogle Scholar
- Henske JK, Wilken SE, Solomon KV, Smallwood CR, Shutthanandan V, Evans JE, Theodorou MK, O'Malley MA. Metabolic characterization of anaerobic fungi provides a path forward for two-stage bioprocessing of crude lignocellulose. Biotechnol Bioeng. 2017. https://doi.org/10.1002/bit.26515 Google Scholar
- Mewis K, Lenfant N, Lombard V, Henrissat B. Dividing the large glycoside hydrolase family 43 into subfamilies: a motivation for detailed enzyme characterization. Appl Environ Microbiol. 2016;82(6):1686–92.View ArticleGoogle Scholar
- Biely P. Microbial carbohydrate esterases deacetylating plant polysaccharides. Biotechnol Adv. 2012;30(6):1575–88.View ArticleGoogle Scholar
- Ali BR, Zhou L, Graves FM, Freedman RB, Black GW, Gilbert HJ, Hazelwood GP. Cellulases and hemicellulases of the anaerobic fungus Piromyces constitute a multiprotein cellulose-binding complex and are encoded by multigene families. FEMS Microbiol Lett. 1995;125(1):15–21.View ArticleGoogle Scholar
- Theodorou MK, Brookman J, Trinci APJ. Anaerobic Fungi. In: Makkar HPS, McSweeney CS, editors. Methods in gut microbial ecology for ruminants. Netherlands: IAEA; 2005. p. 55–6.View ArticleGoogle Scholar
- Solomon KV, Henske JK, Theodorou MK, O’Malley MA. Robust and effective methodologies for cryopreservation and DNA extraction from anaerobic gut fungi. Anaerobe. 2016;38:39–46.View ArticleGoogle Scholar
- Benson DA, Cavanaugh M, Clark K, Karsch-Mizrachi I, Lipman DJ, Ostell J. GenBank. Nucleic Acids Res. 2013;41:D36–D42. https://doi.org/10.1093/nar/gks1195 View ArticleGoogle Scholar
- Tamura K, Stecher G, Peterson D, Filipski A, Kumar S. MEGA6: Molecular Evolutionary Genetics Analysis version 6.0. Mol Biol Evol. 2013;30(12):2725–9.View ArticleGoogle Scholar
- Chenna R. Multiple sequence alignment with the Clustal series of programs. Nucleic Acids Res. 2003;31(13):3497–500.View ArticleGoogle Scholar
- Sievers F, Wilm A, Dineen D, Gibson TJ, Karplus K, Li W, Lopez R, McWilliam H, Remmert M, Soding J, et al. Fast, scalable generation of high-quality protein multiple sequence alignments using clustal omega. Mol Syst Biol. 2011;7:539.View ArticleGoogle Scholar
- Letunic I, Bork P. Interactive tree of life (iTOL) v3: an online tool for the display and annotation of phylogenetic and other trees. Nucleic Acids Res. 2016;44(W1):W242–5.View ArticleGoogle Scholar
- Conesa A, Gotz S, Garcia-Gomez JM, Terol J, Talon M, Robles M. Blast2GO: a universal tool for annotation, visualization and analysis in functional genomics research. Bioinformatics. 2005;21(18):3674–6.View ArticleGoogle Scholar
- Ashburner M, Ball CA, Blake JA, Botstein D, Butler H, Cherry JM, Davis AP, Dolinski K, Dwight SS, Eppig JT, et al. Gene ontology: tool for the unification of biology. The Gene Ontology Consortium. Nat Genet. 2000;25(1):25–9.View ArticleGoogle Scholar
- Bairoch A. The ENZYME database in 2000. Nucleic Acids Res. 2000;28(1):304–5.View ArticleGoogle Scholar
- Li L, Stoeckert CJJ, Roos DS. OrthoMCL: identification of ortholog groups for eukaryotic genomes. Genome Res. 2003;13(9):2178–89.View ArticleGoogle Scholar