MINIREVIEW
Anaerobic fungi (phylum Neocallimastigomycota): advances in
understanding their taxonomy, life cycle, ecology, role and
biotechnological potential
Robert J. Gruninger1, Anil K. Puniya2, Tony M. Callaghan3, Joan E. Edwards3, Noha Youssef4, Sumit
S. Dagar5, Katerina Fliegerova6, Gareth W. Griffith3, Robert Forster1, Adrian Tsang7, Tim McAllister1
& Mostafa S. Elshahed4
1
Agriculture and Agri-Food Canada, Lethbridge, AB, Canada; 2National Dairy Research Institute, Karnal, India; 3IBERS, Aberystwyth University,
Aberystwyth, Wales, UK; 4Oklahoma State University, Stillwater, OK, USA; 5Agharkar Research Institute, Pune, Maharashtra, India; 6Institute of
Animal Physiology and Genetics, Czech Academy of Sciences, Prague, Czech Republic; and 7Concordia University, Montreal, QC, Canada
Correspondence: Mostafa S. Elshahed,
Oklahoma State University, Department of
Microbiology and Molecular Genetics, 1110 S
Innovation Way Dr., Stillwater, OK 74074,
USA. Tel.: +1 (405) 744 3005;
fax: +1 (405) 744 1112;
e-mail: [email protected]
MICROBIOLOGY ECOLOGY
Received 9 June 2014; revised 3 July 2014;
accepted 7 July 2014. Final version published
online 11 August 2014.
DOI: 10.1111/1574-6941.12383
Editor: Gerard Muyzer
Keywords
gut fungi; herbivore; biotechnology; rumen
fungi; fungal genomics.
Abstract
Anaerobic fungi (phylum Neocallimastigomycota) inhabit the gastrointestinal
tract of mammalian herbivores, where they play an important role in the degradation of plant material. The Neocallimastigomycota represent the earliest
diverging lineage of the zoosporic fungi; however, understanding of the relationships of the different taxa (both genera and species) within this phylum is
in need of revision. Issues exist with the current approaches used for their
identification and classification, and recent evidence suggests the presence of
several novel taxa (potential candidate genera) that remain to be characterised.
The life cycle and role of anaerobic fungi has been well characterised in the
rumen, but not elsewhere in the ruminant alimentary tract. Greater understanding of the ‘resistant’ phase(s) of their life cycle is needed, as is study of
their role and significance in other herbivores. Biotechnological application of
anaerobic fungi, and their highly active cellulolytic and hemi-cellulolytic
enzymes, has been a rapidly increasing area of research and development in the
last decade. The move towards understanding of anaerobic fungi using –omics
based (genomic, transcriptomic and proteomic) approaches is starting to yield
valuable insights into the unique cellular processes, evolutionary history, metabolic capabilities and adaptations that exist within the Neocallimastigomycota.
Introduction
Mammalian herbivores do not produce cellulolytic or
hemi-cellulolytic enzymes to degrade ingested plant material; instead they rely on symbiotic associations with
microorganism (i.e. anaerobic fungi, bacteria, methanogenic archaea and protozoa) that reside within their gut.
Within this microbial consortium, anaerobic fungi are
known to be key players in the degradation of lignocellulosic plant fibre in the rumen (Akin et al., 1988, 1989,
1990; Lee et al., 2000a). Zoospores of anaerobic fungi in
the rumen were originally classified as protozoa (Liebetanz, 1910), until it was recognised that these ‘flagellates’
represented the dispersal phase of a zoosporic fungus
(Orpin, 1975). The initial acceptance of these novel
FEMS Microbiol Ecol 90 (2014) 1–17
rumen fungi was complicated; however, due to the longheld belief that all fungi were obligate aerobes. However,
Orpin (1977a) showed that the cell wall of these organisms contained chitin, confirming their correct placement
in kingdom fungi. Since this time, research has started to
uncover the basis for the novel mechanisms that enable
these fungi to live in the absence of oxygen.
Known adaptations of anaerobic fungi to their strict
anaerobic lifestyle include the absence of mitochondria,
cytochromes and other biochemical features of the oxidative phosphorylation pathway (Yarlett et al., 1986; Youssef et al., 2013). Instead, anaerobic fungi possess
specialised organelles called hydrogenosomes, which couple the metabolism of glucose to cellular energy production without the need for oxygen. These organelles have
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R.J. Gruninger et al.
2
features in common with mitochondria (van der Giezen,
2002) and are thought to be derived from them (Embley
et al., 1997, 2003; Voncken et al., 2002; Muller et al.,
2012). Hydrogenosomes contain hydrogenase; producing
H2, CO2, formate and acetate as metabolic waste products
(Brul & Stumm, 1994; Theodorou et al., 1996; Muller
et al., 2012). These, along with lactate and ethanol, are
the main fermentation end products produced from the
anaerobic fungal degradation and fermentation of a
variety of plant cell wall polysaccharides.
Anaerobic fungi degrade recalcitrant lignocellulosic
material both by invasive rhizoidal growth and the associated production of a range of powerful polysaccharidedegrading enzymes. The recently published genome
sequence of Orpinomyces sp. strain C1A has provided
valuable insight into the diversity of these carbohydrate
active enzymes (Youssef et al., 2013), many of which
appear to have been acquired by horizontal gene transfer
from rumen bacteria. The cellulolytic machinery of anaerobic fungi consists of both free enzymes, as well as high
molecular weight extracellular multi-enzyme complexes
(Wilson & Wood, 1992) called cellulosomes (Krause
et al., 2003; Joblin et al., 2010). These potent enzymes
have received much attention in recent years, particularly
in terms of their biotechnological application. This review
covers the key advances that have been made in this area
over the last decade, as well as numerous other areas of
anaerobic fungal research, and highlights the challenges
and opportunities that currently exist within this research
field.
Taxonomy
Anaerobic fungi belong to the phylum Neocallimastigomycota, the earliest diverging lineage unequivocally assigned
to kingdom Fungi and are closely related to the chytrids
(phylum Chytridiomycota) (James et al., 2006a,b; Hibbett
et al., 2007). While they share key morphological features
with their chytrid relatives, they possess a distinctive
anaerobic physiology and flagellar apparatus. Furthermore, genetic analyses have consistently shown that the
Neocallimastigomycota form a distinct, well-supported
clade basal to the chytrids (Fliegerova et al., 2004; James
et al., 2006a; Powell & Letcher, 2012). The phylum Neocallimastigomycota currently comprises six genera, each
distinguishable by morphological features: thallus morphology (rhizoidal vs. bulbous) and zoospore flagellation
(monoflagellate vs. polyflagellate) (Ho & Barr, 1995; Ozkose et al., 2001). These features are summarised in
Table 1, along with some illustrative microscopy images
(Fig. 1). Although conclusive genus identification of
anaerobic fungi using microscopic approaches can be
challenging, assignment of isolates into bulbous (Caecomyces, Cyllamyces), hyphael monocentric (Neocallimastix
and Piromyces) and hyphael polycentric (Orpinomyces and
Anaeromyces) by direct examination of colonies/cultures
is reasonably straightforward (Griffith et al., 2009). However, difficulties in observing zoospore release (Ho &
Bauchop, 1991), the pleomorphic growth form and variable sporangial morphology of some isolates (Ho & Barr,
1995; Brookman et al., 2000a; Leis et al., 2013) can make
further identification challenging. This has led to disagreement as to the validity and distinctiveness of some
taxa (Wubah et al., 1991; Ho & Barr, 1995). Prior to the
advent of DNA barcoding, the difficulty of transporting
viable cultures and the absence of any established repository for these fungi also made interlab comparisons very
difficult.
Application of DNA bar coding has served to highlight
these problems; however, sharing of data via the GenBank
repository has also provided a route towards a more
robust reappraisal of these fungi. Genotypic analysis was
initiated by Dore & Stahl (1991), who used partial
sequence of the ribosomal RNA (rRNA) small subunit
(SSU; 18S; c. 1800 bp in total) to confirm the monophyly
of the anaerobic fungi. However, the 18S region is highly
conserved in Neocallimastigomycota making it difficult to
determine relationships between more closely related taxa
(Dagar et al., 2011). Subsequent genetic classification has
mostly focused on the more variable internal transcribed
spacer (ITS) region of the rRNA locus, now formally
agreed as the primary DNA barcode region for all fungi
(Schoch et al., 2012).
PCR amplification of the ITS region relies on primers
which bind to the highly conserved flanking 18S and 28S
(large subunit; LSU) regions, yielding an amplicon of
Table 1. Morphological classification of the currently recognised anaerobic fungal genera
Genus
Thallus
Rhizoids
Flagella per zoospore
Neocallimastix
Piromyces
Caecomyces
Orpinomyces
Anaeromyces
Cyllamyces
Monocentric
Monocentric
Monocentric
Polycentric
Polycentric
Polycentric
Filamentous
Filamentous
Bulbous
Filamentous
Filamentous
Bulbous
Polyflagellate
Uniflagellate,
Uniflagellate,
Polyflagellate
Uniflagellate
Uniflagellate,
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(7–30)
sometimes bi- or quadriflagellate
sometimes bi- or quadriflagellate
(14–24)
sometimes bi- or triflagellate
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3
Anaerobic fungi: the who, what, why, where and how useful
(a)
(b)
(c)
(d)
(e)
(f)
Fig. 1. Phase contrast (a–d) and bisbenzimide stained fluorescence (e and f) microscopy images of various anaerobic fungal morphological
features (scale bar, 50 lm): (a) free polyflagellated zoospores in a mixed culture; (b) germination and rhizoidal development of a monocentric,
filamentous Piromyces sp.; (c) Polycentric sporangia of an Anaeromyces sp.; (d) bulbous rhizoidal system of Caecomyces sp.; (e) nuclear migration
to rhizoids (white arrow) in a polycentric isolate; (f) nucleated rhizoids of Orpinomyces joyonii.
600–700 bp. The ITS region comprises two type I introns
(ITS1 and ITS2) split by the 5.8S rRNA subunit (159 bp
long), the latter providing an additional region for design
of conserved primers and amplification of ITS1 or ITS2
separately. To date, the ITS1 region has been the more
widely used amplicon for comparison of different genera
and species of anaerobic fungi (Li & Heath, 1992; Brookman et al., 2000a; Fliegerova et al., 2004; Tuckwell et al.,
2005). These analyses consistently support the close relationship between the two genera which form polyflagellate zoospores (Neocallimastix, Orpinomyces) and also the
two genera which form bulbous holdfasts (Caecomyces
and Cyllamyces). However, the phylogenetic relatedness of
the rhizoidal genera with monoflagellate zoospores (Piromyces and Anaeromyces) is less clear, and it seems likely
that the genus Piromyces is polyphyletic and in need of
reappraisal (Brookman et al., 2000a; Hausner et al., 2000;
Fliegerova et al., 2004).
In addition to its application in taxonomic studies, the
ITS1 locus has been widely used in culture-independent
studies to assess fungal diversity and community structure. Various techniques utilising ITS-based PCR amplicons have been employed in such studies, including
DGGE (Kittelmann et al., 2012), ARISA (Edwards et al.,
2008; Cheng et al., 2009; Sundset et al., 2009), clone
library sequencing (Fliegerova et al., 2006; Denman et al.,
2008; Nicholson et al., 2010) and next-generation
FEMS Microbiol Ecol 90 (2014) 1–17
sequencing (Liggenstoffer et al., 2010; Kittelmann et al.,
2012, 2013). Furthermore, ITS1-based approaches have
also been used for the quantification of anaerobic fungi
in environmental samples by qPCR (Denman & McSweeney, 2006; Edwards et al., 2008; Sekhavati et al., 2009;
McDonald et al., 2010; Lwin et al., 2011; Kittelmann
et al., 2012; Marano et al., 2012). The utilisation of these
approaches has clearly demonstrated that the scope of
diversity of anaerobic fungi is significantly wider than
previously implied by culture-based studies (see Ecology).
Despite the widespread use of ITS1 as the formal fungal
barcode (Schoch et al., 2012), it is apparent that its use
for anaerobic fungi is problematic. High sequence variability can cause difficulties in ITS1 sequence alignment,
while intragenomic variation within the ITS region causes
problems for direct sequencing of PCR products (Li &
Heath, 1992; Brookman et al., 2000a; Hausner et al.,
2000; Ozkose et al., 2001; Fliegerova et al., 2004; Nicholson et al., 2005, 2010; Chen et al., 2007; Edwards et al.,
2008). These problems have also been found for some
other groups of fungi (O’Donnell & Cigelnik, 1997; Ko &
Jung, 2002; Nilsson et al., 2008). Lastly, the misidentification of sequences in NCBI database has caused confusion
when attempting to classify novel sequences (Bidartondo,
2008; Kittelmann et al., 2012). To start addressing this
issues, a revised taxonomic framework for the anaerobic
fungi has recently been proposed (Fig. 2; reprinted with
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Published by John Wiley & Sons Ltd. All rights reserved
4
R.J. Gruninger et al.
Fig. 2. Profile Neighbour Joining tree of
Neocallimastigomycota ITS1 sequences based
on 575 unique ITS1 sequences across the
Neocallimsatigomycota. The predicted
secondary structure of the ITS1 region was
modelled to generate an improved alignment
of sequences. In addition to the six named
genera, it is apparent that ten or more clades,
which at present are unnamed, also exist. This
figure has been reprinted with permission
from Koetschan et al. (2014).
permission from Koetschan et al., 2014) in the light of
the large amounts of next-generation sequencing data that
has being generated, and the discovery of many novel
candidate genera that remain to be cultivated (Liggenstoffer et al., 2010; Kittelmann et al., 2012; Koetschan et al.,
2014; see also Ecology). A website has also been created
to allow for researchers to use the secondary structure
prediction approach used by Koetschan et al. (2014) to
incorporate new ITS1 sequences from anaerobic fungi
into this classification scheme (https://www.anaerobicfungi.biocommons.org.nz).
Life cycle
Anaerobic fungi reproduce through the asexual production of flagellated zoospores from sporangia (Heath et al.,
1986), with no sexual reproductive life stage identified to
date. Zoospores may be posteriorly monoflagellate or
polyflagellate, with the subsequently developed thallus
being either monocentric (single reproductive body i.e.
one sporangium from single zoospore) or polycentric
(many sporangia from a single zoospore) (Ho & Barr,
1995). In the rumen, zoospores are released from anaerobic fungal sporangia in response to ingestion of food,
with the timing of peak zoospore density being reached
within 30–60 min (Orpin, 1975, 1976, 1977b; Orpin &
Joblin, 1997). Ruminal zoospore differentiation and subsequent maturation are thought to occur through the
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induction of sporangia by haem and other related porphyrins (Fig. 3), which are released from ingested plant
material (Orpin & Greenwood, 1986).
The locomotion of released Neocallimastix frontalis
zoospores is provided by beating of up to 8–17 flagella
which form a locomotory organelle, the activity of which
has been described previously (Lowe et al., 1987a,b,c).
Motile zoospores locate plant material for colonisation
via chemotactic responses to soluble sugars (Orpin &
Bountiff, 1978) and/or phenolic acids (Wubah & Kim,
1996). Flagellated zoospores of Piromyces communis and
N. frontalis show chemotaxis in vivo towards stomata and
lateral spikes on ingested plant material, with certain soluble sugars leaking from these and other areas of damaged tissues creating a gradient towards which zoospores
are preferentially attracted (Orpin, 1975, 1977b; Orpin &
Bountiff, 1978). The high sensitivity of anaerobic fungal
zoospores to soluble sugars means that freshly ingested
food is quickly colonised, prior to or at the same time as
other rumen microorganisms (Orpin & Bountiff, 1978;
Edwards et al., 2008). As soluble carbohydrates are generally depleted 2–3 h after feeding in sheep, rapid colonisation of plant material by anaerobic fungi is crucial in the
competitive environment of the rumen (Edwards et al.,
2008).
Following attachment to plant material, the zoospores
shed their flagella to form a cyst, although amoeboid
movement across the plant surface has occasionally been
FEMS Microbiol Ecol 90 (2014) 1–17
Anaerobic fungi: the who, what, why, where and how useful
5
Fig. 3. Summary of the anaerobic fungal life
cycle. The stages in the life cycle where
‘resistant’ structures (that have been reported
to date) may be formed are also indicated (*).
observed (Orpin, 1975). Cyst formation and germination
involve the thickening of the cell wall and production of
a germ tube from the polar end opposite from where the
flagella originated (Orpin, 1975, 1994). Cyst development
varies depending on whether the fungus is mono- or
poly-centric (Fig. 3; Table 1). In monocentric taxa, cyst
germination is termed endogenous, as the nucleus
remains within the cyst which enlarges to form a zoosporangium. Thus the rhizoids remain anucleate. In contrast
polycentric taxa exhibit exogenous development, with
migration of nuclei into the more extensive rhizoidal system and thereby enabling the formation of multiple sporangia on each thallus (Trinci et al., 1994; Orpin &
Joblin, 1997). The terminology above is less clear when
applied to the two genera which form bulbous holdfasts
(Caecomyces/Cyllamyces), as discussed by Ozkose et al.
(2001). In both cases nuclei are observed in the holdfast,
consistent with exogenous development, and in the case
of Cyllamyces, also in the branched sporangiophores.
FEMS Microbiol Ecol 90 (2014) 1–17
However, the development of these thalli, while not as
strictly determinate as the monocentric/rhizoidal Neocallimastix and Piromyces, is clearly more limited than the
polycentric/rhizoidal Anaeromyces/Orpinomyces.
The anaerobic fungal rhizomycelium physically penetrates and enzymatically digests plant material while providing an anchor for the production of an external
sporangium (monocentric) or multiple sporangia (polycentric) (Lowe et al., 1987b; Orpin & Joblin, 1997). The
rhizomycelium is categorised as being either filamentous
or bulbous (Table 1, Fig. 1), the latter possessing spherical holdfasts (Ozkose et al., 2001; Chen et al., 2007). The
developing rhizoids are capable of physically penetrating
rigid, undamaged plant cell walls using an appressoriumlike structure (Ho et al., 1988a,b). This process opens up
internal plant tissues to enzymatic breakdown, providing
nutrients that enable the development and maturation of
the multinucleate sporangia (Fig. 3). These mature sporangia may produce a few (1 or 2) to many (50–80)
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6
zoospores (Heath et al., 1983; Lowe et al., 1987a). In the
presence of suitable inducers, the mature sporangium
then undergoes zoospore differentiation, followed by the
release of its zoospores by the dissolution of the sporangial wall.
The fact that it is very difficult to maintain host animals free of anaerobic fungi (Becker, 1929) attests to the
efficient dispersal of anaerobic fungi between hosts, presumably via the formation of aerotolerant survival structures. Several studies have demonstrated that anaerobic
fungi can be cultured from faecal material following airdrying, freezing and even from cow dung left for many
months under field conditions (Lowe et al., 1987c; Milne
et al., 1989; Davies et al., 1993a; McGranaghan et al.,
1999; Griffith et al., 2009). Of the resistant structures that
have been observed to date, probably the most convincing
are the 2–4 chambered spores formed by some Anaeromyces cultures (Brookman et al., 2000b; Ozkose, 2001)
although the processes whereby these form and later germinate are currently unknown.
Ecology
Multiple reports on the isolation of anaerobic fungi from
various herbivores have been published, collectively confirming their detection in 24 different host genera representing eight different animal families (Supporting
Information, Table S1). On the basis of these reports, it
appears that the establishment of anaerobic fungi in the
gut of herbivores has two main prerequisites: a digestive
process involving long resident times for ingested plant
material and a dedicated digestive chamber (e.g. rumen,
forestomach or caecum) with a relatively neutral pH.
Anaerobic fungi appear to be present in all foregut
fermenters (i.e. those where the majority of plant fermentation occurs prior to gastric digestion) including:
ruminants (families Bovidae and Cervidae), pseudoruminants (i.e. those with a three-chambered stomach, e.g.
hippopotamus, camels, llamas, alpaca and vicuna) and
foregut nonruminants (animals possessing an enlarged
forestomach, e.g. marsupials) (Table S1). Anaerobic fungi
have also been detected in multiple hindgut fermenters
(i.e. those where the majority of plant fermentation
occurs postgastric digestion in the caecum and large
intestine, e.g. elephants, horses and rhinoceros) where
they appear to be a normal part of the gut microbial
community (Table S1). They have also been identified in
some larger herbivorous rodents such as the Mara
(Dolichotis patagonum); Teunissen et al., 1991), but not
in other small animals with a hindgut fermentation (presumably due to the shorter duration time of ingested
plant material in their smaller caecum). Interestingly,
anaerobic fungi appear to be absent in strict herbivorous
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R.J. Gruninger et al.
mammals lacking foregut and hindgut fermentation
chambers such as the Panda (Milne et al., 1989), presumably due to the simplicity of their alimentary tract.
Within the reptiles (Class Reptilia), microscopic (Mackie et al., 2004) and molecular identification of anaerobic
fungi (Liggenstoffer et al., 2010) (Table S2) have been
reported in the family Iguanidae; however, they have not
been identified in other herbivorous reptiles (i.e. Tortoises) (Liggenstoffer et al., 2010). The microscopic evidence of structures resembling chytrid thalli (similar to
Caecomyces spp.) and zoospores in the gut of a burrowing
irregular sea urchin (Echinocardium cordatum), however,
is truly intriguing; potentially representing the first documentation of anaerobic fungi in a nonherbivorous (detritivore) marine invertebrate (Thorsen, 1999). The
detection of Neocallimastigomycota DNA in soil (Lockhart
et al., 2006), and estuarine sediments (Devon and Martiny, 2011), is also consistent with their ability to disperse
efficiently between hosts (Life cycle). However, the failure
of most efforts to isolate anaerobic fungi from nongut
habitats (various personal communications; Wubah &
Kim, 1995) suggests that it is only the aerotolerant propagules of these fungi that remain viable in the wider environment.
The geographical distribution of anaerobic fungi is
wide, for example, Cyllamyces (first identified in the UK)
has since been found to be widely distributed in domestic
and wild animals in Africa, Australia, Czech Republic,
India and the USA (Sridhar et al., 2007; Fliegerova et al.,
2010; Liggenstoffer et al., 2010; Nicholson et al., 2010;
Sirohi et al., 2013b). On this basis, it seems more likely
that factors such as host phylogeny, gut type and diet are
more important than geographical location in determining populations of anaerobic fungi in the herbivore gut.
From the broad range of studies performed with domestic
ruminants, it is apparent that diet is one of the key factors affecting this population (Denman et al., 2008; Khejornsart & Wanapat, 2010; Belanche et al., 2012;
Kittelmann et al., 2012; Boots et al., 2013; Sirohi et al.,
2013a), although many of these studies focussed primarily
on quantitation of anaerobic fungi. In contrast to the
effect of diet, systematic studies of the effect of host phylogeny and/or gut type on anaerobic fungi have been
more limited (Liggenstoffer et al., 2010).
Association patterns between specific animal hosts and
anaerobic fungi can be elucidated from the large body of
literature describing the isolation of these microorganism.
Culture-dependent isolation surveys collectively suggest a
pattern in which Piromyces and Caecomyces are the most
prominent genera in the alimentary tract of hindgut fermenters, although the isolation of Anaeromyces from
mules has also been reported (Table S1 and references within). No concrete evidence for the recovery of
FEMS Microbiol Ecol 90 (2014) 1–17
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Anaerobic fungi: the who, what, why, where and how useful
Neocallimastix, Orpinomyces or Cyllamyces from hindgut
fermenters is currently available. While the communities
of anaerobic fungi within the alimentary tract of hindgut
fermenters appear to be dominated by a subset of the
currently described genera, this does not appear to be the
case in ruminants. Ruminal anaerobic fungi are more
diverse, with representatives of all six described genera
having been isolated from domesticated and wild ruminants. Isolates belonging to the genus Neocallimastix
appear to be the most commonly recovered, followed by
members of the genus Piromyces. Orpinomyces and Anaeromyces have also been isolated from several domesticated
and wild ruminants. In contrast, members of the genus
Cyllamyces appear to have a fairly limited host distribution, having only been recovered from domesticated cattle
to date. Finally, although more commonly encountered in
hindgut fermenters, Caecomyces have also been isolated
from domesticated cattle (See Table S2 and references
within). Although there appears to be a distinct distribution of anaerobic fungi within various host animals,
transfer of anaerobic fungi between different animal species in nature appears plausible. Indeed, anaerobic fungi
have been successfully transplanted from horse and reindeer to sheep (Orpin, 1989).
The increasing use of cultivation-independent
approaches in recent years has given additional insight
into the diversity and community structure of anaerobic
fungi in the herbivorous gut and enabled the identification of sixteen different putative novel lineages (Candidate genera) within the Neocallimastigomycota (Table S2).
Liggenstoffer et al. (2010) used a pyrosequencing-based
approach to survey populations of anaerobic fungi in a
large number of diverse zoo animals; concluding that animal host phylogeny appeared to be the most important
factor in determining anaerobic fungal community structure and diversity. The findings of this study are also in
agreement with the observation that Piromyces and Caecomyces are the most prevalent genera cultivated from
hindgut fermenters (Table S1). Interestingly, the majority
of sequences obtained from horses represented novel taxa
(Table S2). Within foregut fermenters, genus-level diversity was generally higher relative to hindgut fermenters,
with Neocallimastix and Piromyces being the most prevalent known genera (Liggenstoffer et al., 2010). The dominance of Neocallimastix and Piromyces has also been seen
in subsequent studies that extensively sampled a narrower
range of foregut herbivores (sheep, deer and cows) over
different seasons (Kittelmann et al., 2012, 2013). Additional studies have found that Orpinomyces can also be
abundant in the foregut (Sirohi et al., 2013b). Similar to
the currently described genera of anaerobic fungi, several
of the novel candidate genera also show evidence of distinct host distribution patterns (Table S2).
FEMS Microbiol Ecol 90 (2014) 1–17
In addition to the ecological insight obtained from
these recent sequence-based surveys, these studies have
made substantial progress in furthering our understanding of the global genus-level diversity that exists within
the Neocallimastigomycota. The existence of new taxa is
perhaps unsurprising (Orpin, 1994; Fliegerova et al.,
2010; Nicholson et al., 2010), although it is unclear
whether these novel candidate genera are uncultivable
using current methods or whether their occurrence has
merely been overlooked due to microscopic resemblance
to known genera. The relative lack of overlap between
novel lineages identified in recent studies is striking
(Fig. 2, Koetschan et al., 2014) and suggests that additional, yet-unknown novel candidate genera may exist in
nature.
Significance for other gut
microorganism and the host
In axenic culture, anaerobic fungi produce a variety of
metabolic end products including acetate, formate, lactate, ethanol, hydrogen and carbon dioxide (Bauchop &
Mountfort, 1981; Cheng et al., 2009). In the rumen, however, this metabolic profile shifts due to interspecies
hydrogen transfer with physically associated methanogens
(Orpin & Joblin, 1997; Voncken et al., 2002), resulting in
the energetically favourable disposal of electrons via
methanogenesis (Cheng et al., 2009). Enhanced anaerobic
fungal enzyme production and fibre dedgradation have
been reported to occur as a consequence of this favourable interaction (Bauchop & Mountfort, 1981; Mountfort
et al., 1982; Teunissen et al., 1992; Cheng et al., 2009);
however, interactions with other rumen microorganism
are not always so mutually beneficial (Gordon & Phillips,
1998). Incubation of anaerobic fungi with protozoa inhibits fungal-mediated plant cell wall degradation (Lee et al.,
2000a); presumably due to protozoal predation of zoospores and potential damage caused to fungal cell walls by
protozoal enzymes (Morgavi et al., 1994; Miltko et al.,
2014). The mechanical and enzymatic degradation of
plant material by anaerobic fungi provide an increased
surface area for bacterial colonisation (Ho et al., 1988a,b;
Orpin & Joblin, 1997), resulting in an increase in the degradation of plant cell walls (Lee et al., 2000a). It has been
reported; however, that some rumen bacteria can have a
negative impact on anaerobic fungi (Gordon & Phillips,
1998).
In contrast to the large number of rumen-based studies, little is known about the role and microbial interactions of anaerobic fungi in the other gut organs of
ruminants. Anaerobic fungi have been found to be present throughout the alimentary tract of cattle and sheep
(Davies et al., 1993b; Rezaeian et al., 2004), with the
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8
quantity of anaerobic fungi highest in the rumen/omasum
and then decreasing exponentially further down the tract
after the abomasum (Davies et al., 1993b). Anaerobic
fungal communities sampled in the rumen, duodenum
and rectum of domesticated cattle have been shown to be
similar (Jimenez et al., 2007), suggesting that the higher
proportion of anaerobic fungi with a resistance phenotype
in the ruminant hindgut (Davies et al., 1993a) is due to
an altered physiological state. The implications that this
has for the functionality of anaerobic fungi in the ruminant hindgut, as well as for nonruminants, are currently
unclear.
In the rumen, anaerobic fungi are considered to contribute significantly to the overall metabolism of their
host by playing a major role in the degradation of lignified plant tissues (Akin et al., 1990). Experiments, where
anaerobic fungi were either absent or eliminated from the
rumen, have provided insight into the contribution of
anaerobic fungi to fibre digestion, feed intake, rumen fermentation and overall rumen metabolism. Removal of
anaerobic fungi from the rumen results in a decrease in
voluntary feed intake and dry matter degradation, indicating digestion of feed is impaired in these animals
(Akin et al., 1989; Morrison et al., 1990; Gordon & Phillips, 1998). Furthermore, the elimination of anaerobic
fungi from the rumen of sheep also significantly reduced
the degradation of dry matter, neutral detergent fibre,
acid detergent fibre and the activity of carboxymethylcellulase (Ford et al., 1987; Gordon & Phillips, 1993; Gao
et al., 2013).
Although the importance of anaerobic fungi in carbohydrate digestion is well established, the role they play in
protein metabolism in the rumen is not clear as the protease activity of anaerobic fungal isolates has been found
to vary significantly (Michel et al., 1993; Yanke et al.,
1993). Anaerobic fungi produce extracellular proteases
that display catalytic activity comparable to the highly
active proteases produced by numerous rumen bacteria,
the majority of which are most active at typical rumen
pH (Wallace & Joblin, 1985; Asao et al., 1993; Bonnemoy
et al., 1993; Michel et al., 1993). Anaerobic fungal proteases provide amino acids for fungal growth, as well as
aiding the penetration of plant material. In addition to
producing enzymes, anaerobic fungi themselves contribute to the protein supply of the host by serving as a
source of high-quality microbial protein, that is synthesised in the rumen, passing to the abomasum and intestines for subsequent digestion and absorption by the host
(Kemp et al., 1985; Gulati et al., 1989; Atasoglu & Wallace, 2002).
In recent years, there has been great interest in the biohydrogenation of lipids in the rumen, particularly the formation of conjugated linoleic acid. Numerous studies
ª 2014 Federation of European Microbiological Societies.
Published by John Wiley & Sons Ltd. All rights reserved
R.J. Gruninger et al.
have investigated the biohydrogenation potential of
anaerobic fungi, with seemingly contrasting findings
reported regarding their enzymatic capabilities and activities (Kemp et al., 1984; Maia et al., 2007; Nam & Garnsworthy, 2007a,b,c). Like proteolysis, however, the rate
and extent of biohydrogenation activity also appears to
vary greatly among anaerobic fungal isolates (Nam &
Garnsworthy, 2007a). In general, anaerobic fungi appear
to play a less active role in biohydrogenation than rumen
bacteria, suggesting that their contribution to this process
is limited. Anaerobic fungi themselves are also unlikely to
serve as a source of polyunsaturated fatty acids to the
host, as N. frontalis only contains saturated (48%) and
monounsaturated (52%) fatty acids (Body & Bauchop,
1985).
Biotechnological applications
The numerous benefits demonstrated from the presence
of anaerobic fungi within the rumen have led to an
increasing interest in their use as a probiotic over the last
decade. The application of anaerobic fungi as a direct-fed
microbial supplement has been investigated, in both
ruminant and nonruminant livestock production, as a
means to improve utilisation of low-quality forages.
Inclusion of cultures of anaerobic fungi in the diets of
various ruminants has been investigated and demonstrated to improve feed intake, animal growth rate, feed
efficiency and milk production (Lee et al., 2000b; Dey
et al., 2004; Paul et al., 2004, 2011; Tripathi et al., 2007;
Samanta et al., 2008; Sehgal et al., 2008; Mamen et al.,
2010; Saxena et al., 2010; Gao et al., 2013). The benefits
observed being even more marked in young ruminants
(Theodorou et al., 1990; Sehgal et al., 2008) and sheep
devoid of anaerobic fungi (Elliott et al., 1987; Gordon &
Phillips, 1993). Collectively these studies illustrate that the
application of anaerobic fungi as a direct-fed microbial
can be used to improve in vivo digestibility by enhancing:
rumen fermentation characteristics (pH, VFA, ammoniaN) rumen microbial populations and cellulolytic enzyme
activities. In contrast, the inclusion of enzymes secreted
by anaerobic fungi alone does not alter rumen fermentation, highlighting the importance of using viable cultures
as a ruminant feed additive (Lee et al., 2000b).
In contrast to the findings with ruminants, anaerobic
fungal enzymes alone appear to be more effective than
viable cultures in monogastric animal (swine and poultry)
production (Theodorou et al., 1996). It is speculated that
this is likely to be due to the inability of anaerobic fungi
to establish in the gastrointestinal tract of these animals.
In swine and poultry diets, cereals containing difficult to
digest nonstarch polysaccharides (b-glucan in barley and
wheat, and arabinoxylans in rye and oats), constitute the
FEMS Microbiol Ecol 90 (2014) 1–17
Anaerobic fungi: the who, what, why, where and how useful
main recalcitrant component of these feedstuffs. These
polymers also have well-known antinutritive effects that
are associated with the propensity of these molecules to
form high molecular weight, viscous aggregates that
reduce intestinal passage rate, decrease diffusion of digestive enzymes, promote endogenous losses and stimulate
unwanted bacterial proliferation (Bedford & Schulze,
1998). Dietary inclusion of anaerobic fungal glycoside
hydrolases (GH) has been found to increase the growth
of broiler chickens by 25%, by aiding the breakdown of
these polymers (Azain et al., 2002).
The most significant challenge for the biotechnological
use of anaerobic fungi, however, as a feed additive or
otherwise, is the difficulty in setting up continuous-flow
cultures for the efficient production of anaerobic fungal
biomass and/or enzymes. For monogastric application,
this has led to attempts to genetically engineer the bacterium Lactobacillus reuteri, a natural component of broiler
gut Microbial communities, to express anaerobic fungal
xylanases and cellulases (Liu et al., 2005a,b, 2007, 2008).
This type of approach, however, is not feasible for ruminant production systems and other applications where
viable anaerobic fungi are needed. Currently, cultivation
of anaerobic fungi requires repeated batch cultures with
frequent transfers that are often difficult to maintain, as
well as being time-consuming and expensive. Processes
using immobilised growing cells seem to be more promising than traditional fermentations with free cells. Studies
examining the immobilisation of monocentric and polycentric fungi have been reported (McCabe et al., 2001,
2003; Nagpal et al., 2009; Sridhar & Kumar, 2010), however, none of these technologies were feasible for commercial use in their current form. The use of anaerobic
fungi as a direct-fed microbial for ruminants will also
require the development of a suitable means of applying
and storing cultures to ensure maintenance of their viability. Progress has already been made identifying a more
suitable strain by isolating a strain of Neocallimastix, that
is more tolerant of oxygen, changes in culture conditions
and requires fewer transfers for maintenance (Leis et al.,
2013). The recent identification of anaerobic fungi in
novel hosts (Ecology) and improved understanding of the
production of resistant states (Life cycle) in the future,
however, may also enable other opportunities to overcome current challenges. Meanwhile, a large amount of
research effort has instead focused specifically on the
exploitation of anaerobic fungal enzymes in various
industries.
Anaerobic fungi produce a broad range of potent polysaccharide-degrading enzymes making them of particular
interest to several industries: brewing, food, textile, paper
and biofuel production. The cellulolytic and hemicellulolytic capacity conferred to the anaerobic fungi by their
FEMS Microbiol Ecol 90 (2014) 1–17
9
plant cell wall-degrading enzymes is greater than some
currently used commercial enzyme mixtures. Extracellular
enzyme preparations from a Piromyces strain and
N. patriciarum have been found to be highly stable and
exhibited a higher capacity to degrade microcrystalline
cellulose than the commercial enzyme products derived
from Trichoderma reesei (Celluclast: cellulase preparation)
and Aspergilus niger (Novozyme: b-glucosidase preparation, Novo-Nordisk, Denmark) (Dijkerman et al., 1997).
In the paper industry, anaerobic fungal cellulases and xylanases provide environmentally friendly methods to treat
paper pulp. As paper processing and pulp bleaching utilises harsh conditions (high temperatures and pH),
rational enzyme engineering using error-prone PCR has
been used to develop a xylanase, derived from Neocallimastix, which is more stable during these processes (Liu
et al., 1999; Wang et al., 2011). Perhaps one of the most
exciting research areas for the application of anaerobic
fungi, however, is that of biofuel production.
There has recently been a push towards developing
renewable fuels from the fermentative production of ethanol from lignocellulosic agricultural residues. This process involves both the breakdown of the lignocellulose
by carbohydrate active enzymes and the conversion of
these hydrolysis products into fermentable sugars. Efforts
to incorporate fibrolytic enzymes from anaerobic fungi
have focused on expressing a range of carbohydrate
active enzymes into a number of aerobic fungal expression strains (Li et al., 2007; Tsai & Huang, 2008; van
Wyk et al., 2010; O’Malley et al., 2012). Currently, the
dominant microbial strain for industrial ethanol production is Saccharomyces cerevisiae, however, the wild-type
strain of this yeast cannot metabolise xylose and arabinose: two important pentoses released during hydrolysis
of hemicellulose. This has focused efforts on genetically
engineering Sacharomyces cerevisiae to facilitate the conversion of xylose to xylulose by incorporating a xylose
isomerase into this organism. Strains of Sacharomyces
cerevisiae expressing xylose isomerase from Piromyces
and/or Orpinomyces have been developed (Kuyper et al.,
2005; van Maris et al., 2007; Madhavan et al., 2009),
and at this time represents the most promising option
for industrial production of ethanol (Bellissimi et al.,
2009). Another approach that has only recently been
investigated is the use of anaerobic fungi to break down
lignocellulose while simultaneously fermenting the resulting sugars to ethanol (Youssef et al., 2013). This
approach was used with the recently sequenced Orpinomyces sp. strain C1A and was found to break down up
to 62.3% of dry weight of corn stover while yielding
0.045–0.096 mg ethanol per mg biomass. Although this
is a relatively minor amount of ethanol, this work shows
that simultaneous saccharification and fermentation is
ª 2014 Federation of European Microbiological Societies.
Published by John Wiley & Sons Ltd. All rights reserved
R.J. Gruninger et al.
10
possible, and future efforts can now be directed towards
enhancing ethanol yield.
A promising source of renewable, environmentally
friendly energy is the production of biogas from the
anaerobic digestion of organic waste. Currently used bioreactors display somewhat low degradation of organic
material (40–60%) (Prochazka et al., 2012) thus, technologies that can improve this efficiency are needed. The
biological pretreatment of crop residues with white and
brown rot fungi has been shown to be effective at
improving biogas production (Ghosh & Bhattacharyya,
1999). However, as biogas production is an anaerobic
process, the inclusion of an aerobic pretreatment step
increases the overall cost of biogas production. In contrast, the direct incorporation of anaerobic fungi into
these bioreactors would eliminate the requirement of an
aerobic predigestion. Incorporation of anaerobic fungi in
bioreactors improved biogas yield for up to 10 days postinoculation, enhancing yield by 4–22% depending on the
substrate and fungal species used (Fliegerova et al., 2012;
Prochazka et al., 2012). Unfortunately, the inability of
anaerobic fungi to survive long term in fermenters, however, makes the application of anaerobic fungi in commercial full-scale biogas production systems unfeasible
using current technologies (Prochazka et al., 2012).
-omic based studies
Anaerobic fungal genomes have an extremely high AT
content (Brownlee, 1989), particularly in their noncoding
regions (Nicholson et al., 2005; Youssef et al., 2013). The
first study to describe genomic sequences from an anaerobic fungus (Orpinomyces sp. OUS1) identified multiple
skeletal genes, secretory pathways, transporters, as well as
numerous genes encoding for central metabolic pathways
(e.g. Pyruvate formate lyase and malate dehydrogenase)
and enzymes for biopolymer degradation (e.g. peptidases
and xylanases) (Nicholson et al., 2005). Since this study,
the development of high throughput sequencing
approaches has provided new tools and opportunities for
sequencing anaerobic fungal genomes, although their
application has been challenging. The use of pyrosequencing technology alone is unfeasible, due to the high adenine–thymine (AT) content of anaerobic fungal genomes
and their prevalence of homopolymeric A and T repeats.
Furthermore, the proliferation of simple sequence repeats
also complicates the assembly of anaerobic fungal
genomes generated using Illumina sequencing technologies.
Using a combination of Illumina Hi-seq and Sanger
sequencing approaches, however, the Joint Genome Institute was the first to generate a draft anaerobic fungal genome. Despite the Piromyces sp. E2 genome being available
online though (http://genome.jgi.doe.gov/PirE2_1/PirE2_
ª 2014 Federation of European Microbiological Societies.
Published by John Wiley & Sons Ltd. All rights reserved
1.home.html), it has not been described in the literature
to date. More recently, Youssef et al. (2013) reported on
the genome sequencing of Orpinomyces sp. strain C1A,
where they used a combined strategy that utilised both
single molecule real time (SMRT) and Illumina sequencing approaches. This recently conceived approach (Koren
et al., 2012) utilises the short read high-accuracy data
obtained by Illumina sequencing to correct errors
encountered in long reads produced by SMRT sequencing. A comparison of the sequencing of the two currently
available anaerobic fungal genomes is shown in Table 2,
although it should be noted that neither of these genomes
have been closed. Despite this, however, genome analysis
has still provided valuable insights into anaerobic fungal
genomic features, metabolic capabilities and cellular processes. In particular, two key factors appear to have
shaped the genome of Orpinomyces sp. strain C1A: its
position as a basal fungal lineage and its unique habitat
within the gut of herbivores.
The position of anaerobic fungi as a basal fungal lineage is reflected in their genome characteristics that are
also present in other early-branching fungal lineages and/
or nonfungal Opisthokonts, but are absent in the Dikarya
(Ascomycetes and Basidiomycetes) genomes. These characteristics include possession of genes indicating the capability for post-translational fucosylation, production of a
complete axoneme and intraflagellar trafficking machinery
proteins, production of a near-complete focal adhesion
machinery, production of a near-complete c-secretase
complex and the production of extracellular protease
inhibitors that have not been previously encountered in
Dikarya. These features are thought to have evolved prior
to fungal separation from an Opisthokonta ancestor and
appear to have subsequently been lost during the evolution of Dikarya (Youssef et al., 2013). In contrast, multiple features observed in the Orpinomyces sp. strain C1A
genome appear to be unique to anaerobic fungi.
Table 2. Comparison of the genome sequencing of Orpinomyces sp.
strain C1A and Piromyces sp. strain E2
Sequencing technologies
Contigs
Scaffolds
Genome size (Mbp)
AT content (%)
Genes predicted
AT content of ORFs (%)
Standard Deviation‡
Piromyces
sp. E2*
Orpinomyces
sp. C1A†
Illumina and Sanger
17 217
1656
71.02
78.1
14 648
70.7
5.1
Illumina and PacBio
32 574
32 574
100.95
83
16 347
73.9
5.8
*http://genome.jgi.doe.gov/PirE2_1/PirE2_1.info.html.
†
http://www.ncbi.nlm.nih.gov/pubmed/23709508.
‡
Standard deviation in open reading frame (ORF) AT content.
FEMS Microbiol Ecol 90 (2014) 1–17
11
Anaerobic fungi: the who, what, why, where and how useful
Many of the unique features of the anaerobic fungal
genome could be considered a consequence of their evolution in the herbivore gut over hundreds of millions of
years. Several genomic features observed in the Orpinomyces sp. strain C1A genome are characteristically associated
with the process of genetic drift; a process that impacts
the genomes of microbial lineages experiencing loweffective population sizes, bottlenecks in vertical transmission and an asexual life style. Genetic drift is characterised by the expansion of genome size, accumulation of
repeats, and gene duplications (Lynch & Conery, 2003;
Kelkar & Ochman, 2012), and an increase in the rate of
nonlethal mutations, which tends to be biased towards
adenine or thymine mutations such as cytosine deamination or guanine oxidation (McCutcheon & Moran, 2012).
Orpinomyces sp. strain C1A has a large genome relative to
other fungal genomes sequenced to date, the presence of
large intergenic regions, high (83.0%) AT content, and a
high level of gene duplication and microsatellite repeats
(Youssef et al., 2013).
In addition to genetic drift, the anaerobic fungal genome shows evidence of multiple adaptations to improve
its fitness in the anaerobic, prokaryote-dominated environment of the herbivore gut. These adaptations include
the dependence on a mixed acid fermentation pathway
for pyruvate metabolism and energy production, the substitution of ergosterol (which requires molecular oxygen
for its biosynthesis) with tetrahymanol and the acquisition of many genes from bacterial donors (Youssef et al.,
2013). Of these adaptations, the latter appears to have
been perhaps most important in improving the plant biomass degradation capacities of this fungus. A large proportion of the genes encoding carbohydrate active
enzymes (CAZYmes) in the Orpinomyces sp. strain C1A
genome originate from known bacterial inhabitants of the
rumen and hindgut of herbivores (Youssef et al., 2013).
This gene acquisition strategy appears to have evolved
Orpinomyces sp. strain C1A from an ancestor with limited
cellulolytic capability to a robust cellulolytic and hemicellulolytic organism (Youssef et al., 2013).
To circumvent the challenges posed by whole genome
assembly, transcriptome based approaches have also
recently been applied to examine the metabolic and functional capacity of the anaerobic fungal genome. Transcriptomics gives an unbiased perspective of gene
transcription in situ providing a truer reflection of metabolic activities than genome-based analysis (Sorek &
Cossart, 2010). The first paper to use this type of
approach employed a combination of transcriptomics and
proteomics techniques to identify carbohydrate active
enzymes that were expressed and secreted by Neocallimastix patriciarum W5 (Wang et al., 2011). Neocallimastix
patriciarum was grown on a number of recalcitrant
FEMS Microbiol Ecol 90 (2014) 1–17
substrates to stimulate expression of cellulases, and the
transcriptome then sequenced using a combination of 454
and Illumina sequencing technologies. A total of 219 glycoside hydrolases from 25 different GH families were
identified, with a number of these enzymes displaying
novel cellulase activities (Wang et al., 2011).
A meta-transcriptome approach has since been used to
examine the activity of rumen eukaryotic microorganism
in Musk-oxen (Ovibos moschatus), through targeted
sequencing of the poly-adenylated mRNA extracted from
rumen solids (Qi et al., 2011). This approach detected
significantly more cellulases (28% of contigs) than that
previously found in a bovine rumen metagenome (8.5%
of contigs) (Qi et al., 2011). The lack of detection of
anaerobic fungal genomic sequences in recent rumen
metagenomic studies is likely a contributing factor to this
observation (Brulc et al., 2009; Hess et al., 2011) and
highlights the need for targeted analysis of anaerobic
fungi when using sequencing based approaches for
‘omics’ analysis.
Conclusions and future directions
The recent application of next-generation sequencing
techniques to identify anaerobic fungi has provided a
great deal of insight into the phylogenetic diversity and
host distribution of these microorganism. The identification of a number of putative uncultured genera should
encourage the development of novel culturing techniques,
in order to attempt to isolate anaerobic fungi representative of these taxa. It is likely that these efforts will further
emphasise the need for an overhaul of the taxonomy of
the Neocallimastigomycota. Increased efforts to understand
the functional diversity of these microorganism are beginning to suggest that our current view of anaerobic fungi
in the rumen is too simplistic. Efforts to study more
anaerobic fungal genera using a combination of ‘-omics’
and traditional techniques will enhance our understanding of their functional diversification which is likely to
have evolved as a means of avoiding niche competition,
as previously proposed (Griffith et al., 2009). Additionally, attempts to genetically engineer and isolate robust
strains of anaerobic fungi will result in wider biotechnological application of anaerobic fungi and/or their
enzymes in the future: making it more feasible to work
with these fungi on a commercial scale and/or in continuous culture.
Acknowledgements
MSE & NY would like to acknowledge financial support
from the US National Science foundation (NSF EPSCoR
award EPS 0814361) and the US Department of
ª 2014 Federation of European Microbiological Societies.
Published by John Wiley & Sons Ltd. All rights reserved
12
Transportation (Sun Grant Initiative award number
DTOS59-07-G-00053). RJG, RF and TAM would like to
acknowledge support from Agriculture and Agri-foods
Canada and Genome Alberta. AT would like to acknowledge support from Genome Canada and Genome Quebec.
TMC would like to gratefully acknowledge funding from
the Aberystwyth Postgraduate Research Studentship. JE
received funding from the Biotechnology and Biological
Sciences Research Council. SSD and AKP would like to
acknowledge funding to visit Aberystwyth University
(UK) from the Stapledon Memorial Trust, UK; DBTCREST award and Network Project of VTCC – Rumen
Microbes (ICAR). KF would like to acknowledge funding
form ‘Ruminomics (project no. 289319 of EC’s 7th
Framework Programme: Food, Agriculture, Fisheries and
Biotechnology)’.
Authors’ contribution
R.J.G., A.K.P., T.M.C., J.E.E. and M.S.E. contributed
equally to this work.
References
Akin D, Borneman W & Windham W (1988) Rumen fungi:
morphological types from Georgia cattle and the attack on
forage cell walls. Biosystems 21: 385–391.
Akin D, Lyon C, Windham W & Rigsby L (1989) Physical
degradation of lignified stem tissues by ruminal fungi. Appl
Environ Microbiol 55: 611–616.
Akin D, Borneman W & Lyon C (1990) Degradation of leaf
blades and stems by monocentric and polycentric isolates of
ruminal fungi. Anim Feed Sci Technol 31: 205–221.
Asao N, Ushida K & Kojima Y (1993) Proteolytic activity of
rumen fungi belonging to the genera Neocallimastix and
Piromyces. Lett Appl Microbiol 16: 247–250.
Atasoglu C & Wallace RJ (2002) De novo synthesis of amino
acids by the ruminal anaerobic fungi, Piromyces communis and
Neocallimastix frontalis. FEMS Microbiol Lett 212: 243–247.
Azain MJ, Li XL, Shah AK & Davies TE (2002) Separation of
the effects of xylanase and b-glucanase addition on
performance of broiler chicks fed barley based diets. Int
Poult Sci Forum 81(Suppl. 1): 111, Abstr. 46.
Bauchop T & Mountfort DO (1981) Cellulose fermentation by
a rumen anaerobic fungus in both the absence and the
presence of rumen methanogens. Appl Environ Microbiol 42:
1103–1110.
Becker ER (1929) Methods of rendering the rumen and
reticulum of ruminants free from their normal infusorian
fauna. Proc Natl Acad Sci USA 15: 435–438.
Bedford MR & Schulze H (1998) Exogenous enzymes for pigs
and poultry. Nutr Res Rev 11: 91–114.
Belanche A, de la Fuente G, Pinloche E, Newbold CJ & Balcells
J (2012) Effect of diet and absence of protozoa on the
ª 2014 Federation of European Microbiological Societies.
Published by John Wiley & Sons Ltd. All rights reserved
R.J. Gruninger et al.
rumen microbial community and on the representativeness
of bacterial fractions used in the determination of microbial
protein synthesis. J Anim Sci 90: 3924–3936.
Bellissimi E, Van Dijken JP, Pronk JT & Van Maris AJ (2009)
Effects of acetic acid on the kinetics of xylose fermentation
by an engineered, xylose-isomerase-based Saccharomyces
cerevisiae strain. FEMS Yeast Res 9: 358–364.
Bidartondo MI (2008) Preserving accuracy in GenBank. Science
319: 1616.
Body DR & Bauchop T (1985) Lipid composition of an
obligately anaerobic fungus Neocallimastix frontalis isolated
from a bovine rumen. Can J Microbiol 31: 463–466.
Bonnemoy F, Fonty G, Michel V & Gouet P (1993) Effect of
anaerobic fungi on the ruminal proteolysis in gnotobiotic
lambs. Reprod Nutr Dev 33: 551–555.
Boots B, Lillis L, Clipson N, Petrie K, Kenny DA, Boland TM
& Doyle E (2013) Responses of anaerobic rumen fungal
diversity (phylum Neocallimastigomycota) to changes in
bovine diet. J Appl Microbiol 114: 626–635.
Brookman J, Mennim G, Trinci A, Theodorou M & Tuckwell
D (2000a) Identification and characterization of anaerobic
gut fungi using molecular methodologies based on
ribosomal ITS1 and 18S rRNA. Microbiology 146: 393–403.
Brookman JL, Ozkose E, Rogers S, Trinci APJ & Theodorou
MK (2000b) Identification of spores in the polycentric
anaerobic gut fungi which enhance their ability to survive.
FEMS Microbiol Ecol 31: 261–267.
Brownlee AG (1989) Remarkably AT-rich genomic DNA from
the anaerobic fungus Neocallimastix. Nucleic Acids Res 17:
1327–1335.
Brul S & Stumm CK (1994) Symbionts and organelles in
ancrobic protozoa and fungi. Trends Ecol Evol 9: 319–324.
Brulc JM, Antonopoulos DA, Miller ME et al. (2009)
Gene-centric metagenomics of the fiber-adherent bovine
rumen microbiome reveals forage specific glycoside
hydrolases. Proc Natl Acad Sci USA 106: 1948–1953.
Chen Y-C, Tsai S-D, Cheng H-L, Chien C-Y, Hu C-Y & Cheng
T-Y (2007) Caecomyces sympodialis sp. nov., a new rumen
fungus isolated from Bos indicus. Mycologia 99: 125–130.
Cheng YF, Edwards JE, Allison GG, Zhu W-Y & Theodorou
MK (2009) Diversity and activity of enriched ruminal
cultures of anaerobic fungi and methanogens grown
together on lignocellulose in consecutive batch culture.
Bioresour Technol 100: 4821–4828.
Dagar SS, Kumar S, Mudgil P, Singh R & Puniya AK (2011)
D1/D2 domain of large-subunit ribosomal DNA for
differentiation of Orpinomyces spp. Appl Environ Microbiol
77: 6722–6725.
Davies DR, Theodorou MK, Brooks AE & Trinci AP (1993a)
Influence of drying on the survival of anaerobic fungi in
rumen digesta and faeces of cattle. FEMS Microbiol Lett 106:
59–63.
Davies DR, Theodorou MK, Lawrence MI & Trinci AP
(1993b) Distribution of anaerobic fungi in the digestive
tract of cattle and their survival in faeces. J Gen Microbiol
139: 1395–1400.
FEMS Microbiol Ecol 90 (2014) 1–17
Anaerobic fungi: the who, what, why, where and how useful
Denman SE & McSweeney CS (2006) Development of a
real-time PCR assay for monitoring anaerobic fungal and
cellulolytic bacterial populations within the rumen. FEMS
Microbiol Ecol 58: 572–582.
Denman S, Nicholson M, Brookman J, Theodorou M &
McSweeney C (2008) Detection and monitoring of
anaerobic rumen fungi using an ARISA method. Lett Appl
Microbiol 47: 492–499.
Devon M & Martiny JBH (2011) Patterns of fungal diversity
and composition along a salinity gradient. ISME J 5: 379–
388.
Dey A, Sehgal JP, Puniya AK & Singh K (2004) Influence of
an anaerobic fungal culture (Orpinomyces sp.)
administration on growth rate, ruminal fermentation and
nutrient digestion in calves. Asian Aus J Anim 17: 820–
824.
Dijkerman R, Bhansing DC, Op den Camp HJ, van der Drift
C & Vogels GD (1997) Degradation of structural
polysaccharides by the plant cell-wall degrading enzyme
system from anaerobic fungi: an application study. Enzyme
Microb Technol 21: 130–136.
Dore J & Stahl D (1991) Phylogeny of anaerobic rumen
Chytridiomycetes inferred from small subunit ribosomal
RNA sequence comparisons. Can J Bot 69: 1964–1971.
Edwards JE, Kingston-Smith AH, Jimenez HR, Huws SA, Skot
KP, Griffith GW, McEwan NR & Theodorou MK (2008)
Dynamics of initial colonization of nonconserved perennial
ryegrass by anaerobic fungi in the bovine rumen. FEMS
Microbiol Ecol 66: 537–545.
Elliott R, Ash AJ, Calderon-Cortes F, Norton BW &
Bauchop T (1987) The influence of anaerobic fungi on
rumen volatile fatty acid concentrations in vivo. J Agric
Sci 109: 13–17.
Embley TM, Horner DA & Hirt RP (1997) Anaerobic
eukaryote evolution: hydrogenosomes as biochemically
modified mitochondria? Trends Ecol Evol 12: 437–441.
Embley TM, van der Giezen M, Horner DS, Dyal PL, Bell S &
Foster PG (2003) Hydrogenosomes, mitochondria and early
eukaryotic evolution. IUBMB Life 55: 387–395.
Fliegerova K, Hodrova B & Voigt K (2004) Classical and
molecular approaches as a powerful tool for the
characterization of rumen polycentric fungi. Folia Microbiol
49: 157–164.
Fliegerova K, Mrazek J & Voigt K (2006) Differentiation of
anaerobic polycentric fungi by rDNA PCR-RFLP. Folia
Microbiol 51: 273–277.
Fliegerova K, Mrazek J, Hoffmann K, Zabranska J & Voigt K
(2010) Diversity of anaerobic fungi within cow manure
determined by ITS1 analysis. Folia Microbiol 55: 319–325.
Fliegerova K, Prochazka J, Mrazek J, Novotna Z, Strosova L &
Dohanyos M (2012) Biogas and rumen fungi. Biogas:
Production, Consumption and Applications (Litonjua R &
Cvetkovski I, eds), pp. 161–179. Nova Science Publishers,
New York, NY.
Ford CW, Elliott R & Maynard PJ (1987) The effect of chlorite
delignification on digestibility of some grass forages and on
FEMS Microbiol Ecol 90 (2014) 1–17
13
intake and rumen microbial activity in sheep fed barley
straw. J Agric Sci 108: 129–136.
Gao AW, Wang HR, Yang JL & Shi CX (2013) The effects of
elimination of fungi on microbial population and fiber
degradation in sheep rumen. Appl Mech Mater 295: 224–231.
Ghosh A & Bhattacharyya BC (1999) Biomethanation of white
rotted and brown rotted rice straw. Bioprocess Eng 20: 297–
302.
Gordon GLR & Phillips MW (1993) Removal of anaerobic
fungi from the rumen of sheep by chemical treatment and
the effect on feed consumption and in vivo fibre digestion.
Lett Appl Microbiol 17: 220–223.
Gordon GLR & Phillips MW (1998) The role of anaerobic gut
fungi in ruminants. Nutr Res Rev 11: 133–168.
Griffith GW, Ozkose E, Theodorou MK & Davies DR (2009)
Diversity of anaerobic fungal populations in cattle revealed
by selective enrichment culture using different carbon
sources. Fungal Ecol 2: 87–97.
Gulati SK, Ashes JR, Gordon GLR, Connell PJ & Rogers PL
(1989) Nutritional availability of amino acids from the
rumen anaerobic fungus Neocallimastix sp. LM1 in sheep. J
Agric Sci 113: 383–387.
Hausner G, Inglis GD, Yanke LJ, Kawchuk LM & McAllister
TA (2000) Analysis of restriction fragment length
polymorphisms in the ribosomal DNA of a selection of
anaerobic chytrids. Can J Bot 78: 917–927.
Heath IB, Bauchop T & Skipp RA (1983) Assignment of the
rumen anaerobe Neocallimastix frontalis to the
Spizellomycetales (Chytridiomycetes) on the basis of its
polyflagellate zoospore ultrastructure. Can J Bot 61: 295–
307.
Heath IB, Kaminskyj SG & Bauchop T (1986) Basal body loss
during fungal zoospore encystment: evidence against
centriole autonomy. J Cell Sci 83: 135–140.
Hess M, Sczyrba A, Egan R et al. (2011) Metagenomic
discovery of biomass-degrading genes and genomes from
cow rumen. Science 331: 463–467.
Hibbett DS, Binder M, Bischoff JF et al. (2007) A higher-level
phylogenetic classification of the Fungi. Mycol Res 111: 509–
547.
Ho YW & Barr DJS (1995) Classification of anaerobic gut
fungi from herbivores with emphasis on rumen fungi from
Malaysia. Mycologia 87: 655–677.
Ho YW & Bauchop T (1991) Morphology of three polycentric
rumen fungi and description of a procedure for the
induction of zoosporogenesis and release of zoospores in
cultures. J Gen Microbiol 137: 213–217.
Ho YW, Abdullah N & Jalaludin S (1988a) Colonization of
guinea grass by anaerobic rumen fungi in swamp buffalo
and cattle. Anim Feed Sci Technol 22: 161–171.
Ho YW, Abdullah N & Jalaludin S (1988b) Penetrating
structures of anaerobic rumen fungi in cattle and swamp
buffalo. J Gen Microbiol 134: 177–181.
James TY, Kauff F, Schoch CL et al. (2006a) Reconstructing
the early evolution of fungi using a six-gene phylogeny.
Nature 443: 818–822.
ª 2014 Federation of European Microbiological Societies.
Published by John Wiley & Sons Ltd. All rights reserved
14
James TY, Letcher J, Longcore SE et al. (2006b) A molecular
phylogeny of the flagellated fungi (Chytridiomycota) and
description of a new phylum (Blastocladiomycota). Mycologia
98: 860–871.
Jimenez HR, Edwards JE, McEwan NR & Theodorou MK
(2007) Characterisation of the population structure of
anaerobic fungi in the ruminant digestive tract. Microb Ecol
Health Dis 19: 40.
Joblin K, Naylor G, Odongo N, Garcia M & Viljoen G (2010)
Ruminal fungi for increasing forage intake and animal
productivity. Sustainable Improvement of Animal Production
and Health (Odongo NE, Garcia M & Viljoen GJ, eds), pp.
129–136. Food and Agriculture Organization of the United
Nations, Rome, Italy.
Kelkar YD & Ochman H (2012) Causes and consequences of
genome expansion in fungi. Genome Biol Evol 4: 13–23.
Kemp P, Lander DJ & Orpin CG (1984) The lipids of the
rumen fungus Piromonas communis. J Gen Microbiol 130:
27–37.
Kemp P, Jordan DJ & Orpin CG (1985) The free- and
protein-amino acids of the rumen phycomycete fungi
Neocallimastix frontalis and Piromonas communis. J Agric Sci
105: 523–526.
Khejornsart P & Wanapat M (2010) Diversity of rumen
anaerobic fungi and methanogenic archaea in swamp buffalo
influenced by various diets. J Anim Vet Adv 9: 3062–3069.
Kittelmann S, Naylor GE, Koolaard JP & Janssen PH (2012) A
proposed taxonomy of anaerobic fungi (Class
Neocallimastigomycetes) suitable for large-scale
sequence-based community structure analysis. PLoS ONE 7:
e36866.
Kittelmann S, Seedorf H, Walters WA, Clemente JC, Knight R,
Gordon JI & Janssen PH (2013) Simultaneous amplicon
sequencing to explore co-occurrence patterns of bacterial,
archaeal and eukaryotic microorganisms in rumen microbial
communities. PLoS ONE 8: e47879.
Ko KS & Jung HS (2002) Three nonorthologous ITS1 types
are present in a polypore fungus Trichaptum abietinum. Mol
Phylogenet Evol 23: 112–122.
Koetschan C, Kittelmann S, Lu J, Al-Halbouni D, Jarvis GN,
M€
uller T, Wolf M & Janssen PH (2014) Internal transcribed
spacer 1 secondary structure analysis reveals a common core
throughout the anaerobic fungi (Neocallimastigomycota).
PLoS ONE 9: e91928.
Koren S, Schatz MC, Walenz BP et al. (2012) Hybrid error
correction and de novo assembly of single-molecule
sequencing reads. Nat Biotechnol 30: 693–700.
Krause DO, Denman SE, Mackie RI, Morrison M, Rae AL,
Attwood GT & McSweeney CS (2003) Opportunities to
improve fiber degradation in the rumen:
microbiology, ecology, and genomics. FEMS Microbiol Rev
27: 663–693.
Kuyper M, Hartog MM, Toirkens MJ, Almering MJ, Winkler
AA, van Dijken JP & Pronk JT (2005) Metabolic
engineering of a xylose-isomerase-expressing Saccharomyces
ª 2014 Federation of European Microbiological Societies.
Published by John Wiley & Sons Ltd. All rights reserved
R.J. Gruninger et al.
cerevisiae strain for rapid anaerobic xylose fermentation.
FEMS Yeast Res 5: 399–409.
Lee SS, Ha JK & Cheng K-J (2000a) Relative contributions of
bacteria, protozoa, and fungi to in vitro degradation of
orchard grass cell walls and their interactions. Appl Environ
Microbiol 66: 3807–3813.
Lee SS, Ha JK & Cheng KJ (2000b) Influence of an anaerobic
fungal culture administration on in vivo ruminal
fermentation and nutrient digestion. Anim Feed Sci Technol
88: 201–217.
Leis S, Dresch P, Peintner U, Fliegerova K, Sandbichler AM,
Insam H & Podmirseg SM (2013) Finding a robust strain
for biomethanation: anaerobic fungi (Neocallimastigomycota)
from the Alpine ibex (Capra ibex) and their associated
methanogens. Anaerobe 29: 34–43.
Li J & Heath IB (1992) The phylogenetic relationships of the
anaerobic chytridiomycetous gut fungi (Neocallimasticaceae)
and the Chytridiomycota. I. Cladistic analysis of rRNA
sequences. Can J Bot 70: 1738–1746.
Li XL, Skory CD, Ximenes EA, Jordan DB, Dien BS,
Hughes SR & Cotta MA (2007) Expression of an AT-rich
xylanase gene from the anaerobic fungus Orpinomyces sp.
strain PC-2 in and secretion of the heterologous enzyme
by Hypocrea jecorina. Appl Microbiol Biotechnol 74:
1264–1275.
Liebetanz E (1910) Die parasitischen Protozoen des
Wiederkauermagen. Arch Protistenkunde 19: 19.
Liggenstoffer AS, Youssef NH, Couger MB & Elshahed MS
(2010) Phylogenetic diversity and community structure of
anaerobic gut fungi (phylum Neocallimastigomycota) in
ruminant and non-ruminant herbivores. ISME J 4: 1225–
1235.
Liu JH, Selinger BL, Tsai C-F & Cheng K-J (1999)
Characterization of a Neocallimastix patriciarum xylanase
gene and its product. Can J Microbiol 45: 970–974.
Liu JR, Yu B, Liu F-H, Cheng K-J & Zhao X (2005a)
Expression of rumen microbial fibrolytic enzyme genes in
probiotic Lactobacillus reuteri. Appl Environ Microbiol 71:
6769–6775.
Liu JR, Yu B, Lin SH, Cheng KJ & Chen YC (2005b) Direct
cloning of a xylanase gene from the mixed genomic DNA of
rumen fungi and its expression in intestinal Lactobacillus
reuteri. FEMS Microbiol Lett 251: 233–241.
Liu JR, Yu B, Zhao X & Cheng KJ (2007) Coexpression of
rumen microbial beta-glucanase and xylanase genes in
Lactobacillus reuteri. Appl Microbiol Biotechnol 77: 117–
124.
Liu JR, Duan C-H, Zhao X, Tzen J, Cheng K-J & Pai C-K
(2008) Cloning of a rumen fungal xylanase gene and
purification of the recombinant enzyme via artificial oil
bodies. Appl Microbiol Biotechnol 79: 225–233.
Lockhart RJ, Van Dyke MI, Beadle IR, Humphreys P &
McCarthy AJ (2006) Molecular biological detection of
anaerobic gut fungi (Neocallimastigales) from landfill sites.
Appl Environ Microbiol 72: 5659–5661.
FEMS Microbiol Ecol 90 (2014) 1–17
Anaerobic fungi: the who, what, why, where and how useful
Lowe SE, Griffith GG, Milne A, Theodorou MK & Trinci APJ
(1987a) The life cycle and growth kinetics of an anaerobic
rumen fungus. J Gen Microbiol 133: 1815–1827.
Lowe SE, Theodorou MK & Trinci AP (1987b) Cellulases and
xylanase of an anaerobic rumen fungus grown on wheat
straw, wheat straw holocellulose, cellulose, and xylan. Appl
Environ Microbiol 53: 1216–1223.
Lowe SE, Theodorou MK & Trinci APJ (1987c) Isolation of
anaerobic fungi from saliva and faeces of sheep. J Gen
Microbiol 133: 1829–1834.
Lwin K, Hayakawa M, Ban-Tokuda T & Matsui H (2011)
Real-time pcr assays for monitoring anaerobic fungal
biomass and population size in the rumen. Curr Microbiol
62: 1147–1151.
Lynch M & Conery JS (2003) The origins of genome
complexity. Science 302: 1401–1404.
Mackie RI, Rycyk M, Ruemmler RL, Aminov RI & Wikelski M
(2004) Biochemical and microbiological evidence for
fermentative digestion in free-living land iguanas
(Conolophus pallidus) and marine iguanas (Amblyrhynchus
cristatus) on the Galapagos archipelago. Physiol Biochem
Zool 77: 127–138.
Madhavan A, Tamalampudi S, Ushida K, Kanai D, Katahira S,
Srivastava A, Fukuda H, Bisaria V & Kondo A (2009)
Xylose isomerase from polycentric fungus Orpinomyces: gene
sequencing, cloning, and expression in Saccharomyces
cerevisiae for bioconversion of xylose to ethanol. Appl
Microbiol Biotechnol 82: 1067–1078.
Maia MRG, Chaudhary LC, Figueres L & Wallace RJ (2007)
Metabolism of polyunsaturated fatty acids and their toxicity
to the microflora of the rumen. Antonie Van Leeuwenhoek
91: 303–314.
Mamen D, Vadivel V, Pugalenthi M & Parimelazhagan T
(2010) Evaluation of fibrolytic activity of two different
anaerobic rumen fungal isolates for their utilization as
microbial feed additive. Anim Nutr Feed Technol 10: 37–
49.
Marano AV, Gleason FH, B€arlocher F et al. (2012)
Quantitative methods for the analysis of zoosporic fungi. J
Microbiol Methods 89: 22–32.
McCabe BK, Kuek C, Gordon GL & Phillips MW (2001)
Immobilization of monocentric and polycentric types of
anaerobic chytrid fungi in Ca-alginate. Enzyme Microb
Technol 29: 144–149.
McCabe BK, Kuek C, Gordon GL & Phillips MW (2003)
Production of b-glucosidase using immobilised Piromyces
sp. KSX1 and Orpinomyces sp. 478P1 in repeat-batch
culture. J Ind Microbiol Biotechnol 30: 205–209.
McCutcheon JP & Moran NA (2012) Extreme genome
reduction in symbiotic bacteria. Nat Rev Microbiol 10: 13–
26.
McDonald JE, Allison HE & McCarthy AJ (2010) Composition
of the landfill microbial community as determined by
application of domain- and group-specific 16S and 18S
rRNA-targeted oligonucleotide probes. Appl Environ
Microbiol 76: 1301–1306.
FEMS Microbiol Ecol 90 (2014) 1–17
15
McGranaghan P, Davies JC, Griffith GW, Davies DR &
Theodorou MK (1999) The survival of anaerobic fungi in
cattle faeces. FEMS Microbiol Ecol 29: 293–300.
Michel V, Fonty G, Millet L, Bonnemoy F & Gouet P (1993)
In vitro study of the proteolytic activity of rumen anaerobic
fungi. FEMS Microbiol Lett 110: 5–9.
Milne A, Theodorou MK, Jordan MGC, King-Spooner C &
Trinci APJ (1989) Survival of anaerobic fungi in feces, in
saliva, and in pure culture. Exp Mycol 13: 27–37.
Miltko R, Belzecki G, Kowalik B & Michalowski T (2014) Can
fungal zoospores be the source of energy for the rumen
protozoa Eudiplodinium maggii? Anaerobe 29: 68–72.
Morgavi DP, Sakurada M, Mizokami M, Tomita Y & Onodera
R (1994) Effects of ruminal protozoa on cellulose
degradation and the growth of an anaerobic ruminal fungus,
Piromyces sp. strain OTS1, in vitro. Appl Environ Microbiol
60: 3718–3723.
Morrison M, Murray RM & Boniface AN (1990) Nutrient
metabolism and rumen micro-organisms in sheep fed a
poor-quality tropical grass hay supplemented with sulphate.
J Agric Sci 115: 269–275.
Mountfort DO, Asher RA & Bauchop T (1982) Fermentation
of cellulose to methane and carbon dioxide by a rumen
anaerobic fungus in a triculture with Methanobrevibacter sp.
strain RA1 and Methanosarcina barkeri. Appl Environ
Microbiol 44: 128–134.
Muller M, Mentel M, van Hellemond JJ, Henze K, Woehle C,
Gould SB, Yu RY, van der Giezen M, Tielens AG & Martin
WF (2012) Biochemistry and evolution of anaerobic energy
metabolism in eukaryotes. Microbiol Mol Biol Rev 76: 444–
495.
Nagpal R, Puniya AK & Singh K (2009) In vitro fibrolytic
activity of the anaerobic fungus, Caecomyces sp.,
immobilized in alginate beads. J Anim Feed Sci 18:
758–768.
Nam IS & Garnsworthy PC (2007a) Biohydrogenation of
linoleic acid by rumen fungi compared with rumen bacteria.
J Appl Microbiol 103: 551–556.
Nam IS & Garnsworthy PC (2007b) Biohydrogenation
pathways for linoleic and linolenic acids by Orpinomyces
rumen fungus. Asian Aust J Anim 20: 1694–1698.
Nam IS & Garnsworthy PC (2007c) Factors influencing
biohydrogenation and conjugated linoleic acid production
by mixed rumen fungi. J Microbiol 45: 199–204.
Nicholson MJ, Theodorou MK & Brookman JL (2005)
Molecular analysis of the anaerobic rumen fungus
Orpinomyces – insights into an AT-rich genome.
Microbiology 151: 121–133.
Nicholson MJ, McSweeney CS, Mackie RI, Brookman JL &
Theodorou MK (2010) Diversity of anaerobic gut fungal
populations analysed using ribosomal ITS1 sequences in
faeces of wild and domesticated herbivores. Anaerobe 16:
66–73.
Nilsson RH, Kristiansson E, Ryberg M, Hallenberg N &
Larsson KH (2008) Intraspecific ITS variability in the
kingdom fungi as expressed in the international sequence
ª 2014 Federation of European Microbiological Societies.
Published by John Wiley & Sons Ltd. All rights reserved
16
databases and its implications for molecular species
identification. Evol Bioinform 4: 193–201.
O’Donnell K & Cigelnik E (1997) Two divergent intragenomic
rDNA ITS2 types within a monophyletic lineage of the
fungus Fusarium are nonorthologous. Mol Phylogenet Evol 7:
103–116.
O’Malley MA, Theodorou MK & Kaiser CA (2012) Evaluating
expression and catalytic activity of anaerobic fungal
fibrolytic enzymes native to Piromyces sp. E2 in
Saccharomyces cerevisiae. Environ Prog Sustain Energy 31:
37–46.
Orpin CG (1975) Studies on the rumen flagellate
Neocallimastix frontalis. J Gen Microbiol 91: 249–262.
Orpin C (1976) Studies on the rumen flagellate Sphaeromonas
communis. J Gen Microbiol 94: 270–280.
Orpin C (1977a) The occurrence of chitin in the cell walls of
the rumen organisms Neocallimastix frontalis, Piromonas
communis and Sphaeromonas communis. J Gen Microbiol 99:
215–218.
Orpin CG (1977b) The rumen flagellate Piromonas communis:
its life-history and Invasion of plant material in the rumen.
J Gen Microbiol 99: 107–117.
Orpin CG (1989) Ecology of rumen anaerobic fungi in relation
to the nutrition of the host animal. The Roles of Protozoa
and Fungi in Ruminant Digestion (Nolan JV, Leng RA &
Demeyer DI, eds), pp. 29–37. Penambul Books, Armidale,
NSW, Australia.
Orpin CG (1994) Anaerobic fungi: taxonomy, biology,and
distribution in nature. Anaerobic Fungi: Biology, Ecology, and
Function (Mountfort DO & Orpin CG, eds), pp. 1–45.
Marcel Dekker Inc., New York, NY, USA.
Orpin CG & Bountiff L (1978) Zoospore chemotaxis in the
rumen phycomycete Neocallimastix frontalis. J Gen Microbiol
104: 113–122.
Orpin CG & Greenwood Y (1986) The role of haems and
related compounds in the nutrition and zoosporogenesis of
the rumen chytridiomycete Neocallimastix frontalis H8. J
Gen Microbiol 132: 2179–2185.
Orpin CG & Joblin KN (1997) The rumen anaerobic fungi.
The Rumen Microbial Ecosystem (Hobson PN & Stewart CS,
eds), pp. 140–184. Blackie Academic and Professional,
London.
Ozkose E (2001) Morphology and Molecular Ecology of
Anaerobic Fungi. University of Wales, Aberyswyth.
Ozkose E, Thomas BJ, Davies DR, Griffith GW & Theodorou
MK (2001) Cyllamyces aberensis gen.nov. sp.nov., a new
anaerobic gut fungus with branched sporangiophores
isolated from cattle. Can J Bot 79: 666–673.
Paul SS, Kamra DN, Sastry VRB, Sahu NP & Agarwal N (2004)
Effect of administration of an anaerobic gut fungus isolated
from wild blue bull (Boselaphus tragocamelus) to buffaloes
(Bubalus bubalis) on in vivo ruminal fermentation and
digestion of nutrients. Anim Feed Sci Technol 115: 143–157.
Paul SS, Deb SM, Punia BS, Das KS, Singh G, Ashar MN &
Kumar R (2011) Effect of feeding isolates of anaerobic
fungus Neocallimastix sp. CF 17 on growth rate and
ª 2014 Federation of European Microbiological Societies.
Published by John Wiley & Sons Ltd. All rights reserved
R.J. Gruninger et al.
fibre digestion in buffalo calves. Arch Anim Nutr 65:
215–228.
Powell MJ & Letcher PM (2012) From zoospores to molecules:
the evolution and systematics of Chytridiomycota.
Systematics and Evolution of Fungi (Progress in Mycological
Research) (Misra JK, Tewari JP & Deshmukh SK, eds), pp.
29–54. CRC Press, New York, NY.
Prochazka J, Mrazek J, Strosova L, Fliegerova K, Zabranska J &
Dohanyos M (2012) Enhanced biogas yield from energy
crops with rumen anaerobic fungi. Eng Life Sci 12:
343–351.
Qi M, Wang P, O’Toole N et al. (2011) Snapshot of the
eukaryotic gene expression in muskoxen rumen—a
metatranscriptomic approach. PLoS ONE 6: e20521.
Rezaeian M, Beakes GW & Parker DS (2004) Distribution and
estimation of anaerobic zoosporic fungi along the digestive
tracts of sheep. Mycol Res 108: 1227–1233.
Samanta AK, Singh KK, Das MM & Pailan GH (2008) Effect
of direct fed anaerobic fungal culture on rumen
fermentation, nutrient utilization and live weight gain in
crossbred heifers. Indian J Anim Sci 78: 1134–1137.
Saxena S, Sehgal JP, Puniya AK & Singh K (2010) Effect of
administration of rumen fungi on production performance
of lactating buffaloes. Benef Microbes 1: 183–188.
Schoch CL, Seifert KA, Huhndorf S, Robert V, Spouge JL,
Levesque CA, Chen W, Bolchacova E, Voigt K & Crous PW
(2012) Nuclear ribosomal internal transcribed spacer (ITS)
region as a universal DNA barcode marker for Fungi. Proc
Natl Acad Sci USA 109: 6241–6246.
Sehgal JP, Jit D, Puniya AK & Singh K (2008) Influence of
anaerobic fungal administration on growth, rumen
fermentation and nutrient digestion in female buffalo calves.
J Anim Feed Sci 17: 510–518.
Sekhavati MH, Mesgaran MD, Nassiri MR, Mohammadabadi
T, Rezaii F & Fani Maleki A (2009) Development and use of
quantitative competitive PCR assays for relative quantifying
rumen anaerobic fungal populations in both in vitro and in
vivo systems. Mycol Res 113: 1146–1153.
Sirohi S, Choudhury P, Dagar S, Puniya A & Singh D (2013a)
Isolation, characterization and fibre degradation potential of
anaerobic rumen fungi from cattle. Ann Microbiol 63: 1187–
1194.
Sirohi S, Choudhury P, Puniya A, Singh D, Dagar S & Singh
N (2013b) Ribosomal ITS1 sequence-based diversity analysis
of anaerobic rumen fungi in cattle fed on high fiber diet.
Ann Microbiol 63: 1571–1577.
Sorek R & Cossart P (2010) Prokaryotic transcriptomics: a
new view on regulation, physiology and pathogenicity. Nat
Rev Genet 11: 9–16.
Sridhar M & Kumar D (2010) Production of fibrolytic enzymes
in repeat-batch culture using immobilized zoospores of
anaerobic rumen fungi. Ind J Biotechnol 9: 87–95.
Sridhar M, Kumar M, Anandan S, Prasad CS & Sampath KT
(2007) Occurrence and prevalence of Cyllamyces genus–a
putative anaerobic gut fungus in Indian cattle and buffaloes.
Curr Sci 92: 1356–1358.
FEMS Microbiol Ecol 90 (2014) 1–17
17
Anaerobic fungi: the who, what, why, where and how useful
Sundset MA, Edwards JE, Cheng YF, Senosiain RS, Fraile MN,
Northwood KS, Praesteng KE, Glad T, Mathiesen SD &
Wright AD (2009) Rumen microbial diversity in Svalbard
reindeer, with particular emphasis on methanogenic
archaea. FEMS Microbiol Ecol 70: 553–562.
Teunissen MJ, Op den Camp HJ, Orpin CG, Huis in ‘t Veld
JH & Vogels GD (1991) Comparison of growth
characteristics of anaerobic fungi isolated from
ruminant and non-ruminant herbivores during
cultivation in a defined medium. J Gen Microbiol 137:
1401–1408.
Teunissen MJ, Kets EP, Op den Camp HJ, Huis in’t Veld JH
& Vogels GD (1992) Effect of coculture of anaerobic fungi
isolated from ruminants and non-ruminants with
methanogenic bacteria on cellulolytic and xylanolytic
enzyme activities. Arch Microbiol 157: 176–182.
Theodorou MK, Beever DE, Haines MJ & Brooks A (1990)
The effect of a fungal probiotic on intake and performance
of early weaned calves. Anim Prod 50: 577.
Theodorou MK, Mennim G, Davies DR, Zhu WY, Trinci AP
& Brookman JL (1996) Anaerobic fungi in the digestive
tract of mammalian herbivores and their potential for
exploitation. Proc Nutr Soc 55: 913–926.
Thorsen MS (1999) Abundance and biomass of the gut-living
microorganisms (bacteria, protozoa and fungi) in the
irregular sea urchin Echinocardium cordatum (Spatangoida:
Echinodermata). Mar Biol 133: 353–360.
Trinci AP, Davies DR, Gull K, Lawrence MI, Bonde Nielsen B,
Rickers A & Theodorou MK (1994) Anaerobic fungi in
herbivorous animals. Mycol Res 98: 129–152.
Tripathi VK, Sehgal JP, Puniya AK & Singh K (2007) Effect of
administration of anaerobic fungi isolated from cattle and
wild blue bull (Boselaphus tragocamelus) on growth rate and
fibre utilization in buffalo calves. Arch Anim Nutr 61: 416–
423.
Tsai CT & Huang C-T (2008) Overexpression of the
Neocallimastix frontalis xylanase gene in the methylotrophic
yeasts Pichia pastoris and Pichia methanolica. Enzyme Microb
Technol 42: 459–465.
Tuckwell DS, Nicholson MJ, McSweeney CS, Theodorou MK
& Brookman JL (2005) The rapid assignment of ruminal
fungi to presumptive genera using ITS1 and ITS2 RNA
secondary structures to produce group-specific fingerprints.
Microbiology 151: 1557–1567.
van der Giezen M (2002) Strange fungi with even stranger
insides. Mycologist 16: 129–131.
van Maris AJ, Winkler AA, Kuyper M, de Laat WT, van
Dijken JP & Pronk JT (2007) Development of efficient
xylose fermentation in Saccharomyces cerevisiae: xylose
isomerase as a key component. Adv Biochem Eng Biotechnol
108: 179–204.
van Wyk N, Den Haan R & van Zyl W (2010) Heterologous
production of NpCel6A from Neocallimastix patriciarum
FEMS Microbiol Ecol 90 (2014) 1–17
in Saccharomyces cerevisiae. Enzyme Microb Technol 46:
378–383.
Voncken F, Boxma B, Tjaden J et al. (2002) Multiple origins
of hydrogenosomes: functional and phylogenetic evidence
from the ADP/ATP carrier of the anaerobic chytrid
Neocallimastix sp. Mol Microbiol 44: 1441–1454.
Wallace RJ & Joblin KN (1985) Proteolytic activity of a rumen
anaerobic fungus. FEMS Microbiol Lett 29: 19–25.
Wang TY, Chen HL, Lu MY et al. (2011) Functional
characterization of cellulases identified from the cow rumen
fungus Neocallimastix patriciarum W5 by transcriptomic and
secretomic analyses. Biotechnol Biofuels 4: 24.
Wilson CA & Wood TM (1992) The anaerobic fungus
Neocallimastix frontalis: isolation and properties of a
cellulosome-type enzyme fraction with the capacity to
solubilize hydrogen-bond-ordered cellulose. Appl Microbiol
Biotechnol 37: 125–129.
Wubah DA & Kim D (1995) Isolation and characterization of
a free-living species of Piromyces from a pond. Inoculum
(Newsletter of the Mycological Society of America; Abstracts
of papers and posters presented at the MSA annual meeting
held 6-10 August, 1995, at the Town and Country Hotel,
San Diego, CA) 46: 52.
Wubah DA & Kim DS (1996) Chemoattraction of anaerobic
ruminal fungi zoospores to selected phenolic acids.
Microbiol Res 151: 257–262.
Wubah DA, Fuller MS & Akin DE (1991) Neocallimastix: a
comparative morphological study. Can J Bot 69: 835–843.
Yanke LJDY, McAllister TA, Bae HD & Cheng KJ (1993)
Comparison of amylolytic and proteolytic activities of ruminal
fungi grown on cereal grains. Can J Microbiol 39: 817–820.
Yarlett N, Orpin CG, Munn EA, Yarlett NC & Greenwood CA
(1986) Hydrogenosomes in the rumen fungus Neocallimastix
patriciarum. Biochem J 236: 729–739.
Youssef NH, Couger MB, Struchtemeyer CG, Liggenstoffer AS,
Prade RA, Najar FZ, Atiyeh HK, Wilkins MR & Elshahed
MS (2013) The genome of the anaerobic fungus
Orpinomyces sp. strain C1A reveals the unique evolutionary
history of a remarkable plant biomass degrader. Appl
Environ Microbiol 79: 4620–4634.
Supporting Information
Additional Supporting Information may be found in the
online version of this article:
Table S1. Anaerobic fungi detected using enrichment and
isolation-based approaches.
Table S2. Culture-independent studies identifying anaerobic fungal diversity and community structure in gut ecosystems.
ª 2014 Federation of European Microbiological Societies.
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