Fungal endophytes: an untapped source of
biocatalysts
Trichur S. Suryanarayanan, Nagamani
Thirunavukkarasu, Meenavalli
B. Govindarajulu & Venkat Gopalan
Fungal Diversity
An International Journal of Mycology
ISSN 1560-2745
Volume 54
Number 1
Fungal Diversity (2012) 54:19-30
DOI 10.1007/s13225-012-0168-7
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Author's personal copy
Fungal Diversity (2012) 54:19–30
DOI 10.1007/s13225-012-0168-7
REVIEW
Fungal endophytes: an untapped source of biocatalysts
Trichur S. Suryanarayanan &
Nagamani Thirunavukkarasu &
Meenavalli B. Govindarajulu & Venkat Gopalan
Received: 27 March 2012 / Accepted: 30 March 2012 / Published online: 18 April 2012
# The Mushroom Research Foundation 2012
Abstract Horizontally transmitted endophytes are an ecological group of fungi that infect living plant tissues and survive in
them without causing any disease symptoms. Even as facets of
the endophyte-plant symbiotic relationship are being uncovered, there is an increasing appreciation of the different growth
substrates exploited by endophytes and the vast repertoire of
secreted enzymes of these fungi. These attributes exemplify
the striking biodiversity of fungal endophytes and should
motivate bioprospecting these organisms to identify novel
biocatalysts that might help address challenges in medicine,
food security, energy production and environmental quality.
Keywords Fungal enzymes . Microbial bioprospecting .
Biodiversity
Introduction
The absorptive mode of nutrition in fungi has resulted in the
evolution and secretion of a battery of enzymes that catabolize complex organic polymers (e.g., cellulose, chitin, protein) in the environment to smaller constituents, which
T. S. Suryanarayanan (*) : M. B. Govindarajulu
Vivekananda Institute of Tropical Mycology (VINSTROM),
RKM Vidyapith,
Chennai 600 004, India
e-mail: [email protected]
N. Thirunavukkarasu
Department of Plant Biology and Plant Biotechnology,
RKM Vivekananda College,
Chennai 600 004, India
T. S. Suryanarayanan : V. Gopalan
Department of Biochemistry and Center for RNA Biology,
The Ohio State University,
Columbus, OH 43210, USA
are then absorbed by their cells for metabolism. That
these polymers need not be broken down to monomeric
units is borne out by the expression in some ascomycete
and basidiomycete fungi of an oligosaccharide transporter
that might play a role in uptake of sucrose (a disaccharide)
and raffinose (a trisaccharide) (Fang and Leger 2010). This
mode of nutrition and the diverse nature of substrates that
the different ecological groups of fungi exploit for growth
help rationalize their ability to employ a diverse array of
extracellular enzymes (e.g., amylases, cellulases, chitinases,
lipases, and proteases). It is therefore not surprising that
around 60 % of the currently used industrial enzymes are
of fungal origin (Østergaard and Olsen 2010); applications
include baking, fermenting coffee beans, processing meat,
manufacturing corn syrup, hydrolyzing milk protein, removing stains, dehairing hides, separating racemic mixtures of
amino acids, biosensing and bioremediation (Table 1).
Although fungi are extraordinarily species rich with
about 1.5 million estimated members (Hawksworth 1991),
merely five genera (Aspergillus, Humicola, Penicillium,
Rhizopus and Trichoderma) account for three quarters of
the 60 % fungal enzymes used in industrial processes
(Østergaard and Olsen 2010) lending immediacy to screening fungi of different ecological groups for novel and more
efficient biocatalysts (Peterson et al. 2011). It is in this
context that we focus here on the need to study endophytes.
Bioprospecting fungal endophytes for novel catalysts:
rationale
Fungal endophytes, an integral part of the plant microbiome,
infect and reside in plants (algae, bryophytes, pteridophytes,
gymnosperms and angiosperms) without initiating any visible disease symptoms (Hyde and Soytong 2008). Although
not universal, infection by endophytes could confer fitness
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Fungal Diversity (2012) 54:19–30
Table 1 Industrial applications of selected enzymes for which endophytes were screened in our studies
Enzyme
Application
Reference
Chitin deacetylase
Zhao et al. 2010
Acidic protease
Preparation of chitosan from chitin; biological control of insect pests, human and plant
pathogens
Drugs for treating asthma, cancer, diabetes, inflammation, wounds, bacterial/fungal
infections; immunity modulation; sialic acid production, anti inflammatory, cosmetics;
waste water treatment
Mediation of drug release; denture cleaners; cosmetics; production of protein hydrolysates;
brewing; baking; animal feeds; waste water treatment; silk degumming; detergent additive;
tanning processes; scavenging silver from x-ray film
Digestive aid; preparation of fermented foods; seasoning material
Tannase
Asparaginase
Clarifying agent in instant tea, wine, fruit juices
Drug for treating acute lymphoblastic leukaemia
Ramírez-Coronel et al. 2003
Schrey et al. 2010
Laccase
Processing of wine, fruit juice and beer; baking; delignification; bio-remediation of
phenolic compounds; bio-bleaching
Biofuel production from lignocellulosic biomass
Kunamneni et al. 2008
Chitinase and
chitosanase
Alkaline protease
β-glucosidase
benefits to the plant host such as tolerance/resistance to
drought, heat, herbivory and disease (Saikkonen et al. 2010;
Vesterlund et al. 2011; Hubbard et al. 2012). Some of the early
focus was on the use of endophytes as biocontrol agents
(Arnold et al. 2003; Vega 2008; Rocha et al. 2011) and as a
possible source of novel bioactive compounds (Gunatilaka
2006; Weber 2009; Aly et al. 2010, 2011; Debbab et al.
2011; Xu et al. 2010). Although endophytes have been studied
rigorously for their secondary metabolites, similar efforts have
not been invested to examine their catalytic repertoire, which
is likely to be vast as suggested by our preliminary investigations on tropical endophytic fungi.
Using agar plate assays for qualitative screening
(Kumaresan et al. 2002), we found that many of the foliar
fungal endophytes from trees of different forests in the
Western Ghats mountain ranges (in southern India) produced
extracellular enzymes including amlyases, cellulases,
chitinases, chitosonases, laccases, lipases, pectinases,
and proteases (Table 2). Such an arsenal of biocatalysts is
not unexpected since endophytes (i) infect plant tissues such
Table 2 Extracellular enzymes produced by endophytes isolated from
trees of tropical forests of the Western Ghats mountains in India
Enzyme
assayed
Number of fungal endophyte
isolates tested
Positive for
activity (%)
Amylase
Cellulase
Laccase
Lipase
Pectinase
Pectate lyase
Protease
Tyrosinase
133
134
134
113
118
134
134
134
60
61
65
84
58
56
59
38
Hartl et al. 2012
Gupta et al. 2002
Rao et al. 1998
Chauve et al. 2010
as leaves, (ii) survive in them and (iii) a few of them continue
to survive in senesced leaves as saprotrophs, thus functioning
as pioneer litter degraders (Kumaresan and Suryanarayanan
2002; Promputtha et al. 2010; Chaverri and Gazis 2011;
Purahong and Hyde 2011). Given this life style, fungal endophytes have to necessarily elaborate a variety of enzymes to
breach the barriers of plants (to infect), to counter the defense
chemicals of plants (to survive in live tissues) and to degrade
the cell walls of senescent plant tissues (to continue as saprotrophs) (Sun et al. 2011). Such versatility provides the underpinning for our expectation that fungal endophyte enzymes
will likely find numerous applications in the realms of health,
food production, energy and environment.
Advances in genomics and proteomics, coupled with highthroughput screening assays, will eventually establish a representative inventory of biocatalysts in all three domains of
life; such a compilation will enable customizing enzymes for
specific applications, inspired by unique biochemical properties of a given biocatalyst. While there is no reason a priori to
expect enzymes from endophytes to be intrinsically superior
catalysts compared to homologs elsewhere, we highlight a
few instances where fungal endophytes might furnish appealing solutions to challenging problems. In the concluding
section, we revisit this theme to provide a roadmap for further
explorations of fungal endophytes.
Applications of enzymes from fungal endophytes
Health and well-being
L-asparaginase
Bacterial L-asparaginase, which hydrolyzes L-asparagine
(Asn) into aspartic acid and ammonia, is used as part of a
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Fungal Diversity (2012) 54:19–30
combinatorial therapy to treat acute lymphoblastic leukemia
(ALL), the most frequent form of cancer among children
(Azevedo-Silva et al. 2010). Because they lack L-asparagine
synthetase, which amidates L-aspartic acid to L-Asn, the
survival of neoplastic lymphoblasts depends on import of LAsn from the blood. Treatment with L-asparaginase depletes
L-Asn in the blood thus starving the malignant cells of L-Asn
leading to protein synthesis defects and ultimately cell death.
Albeit part of successful anti-leukemic approaches, two side
effects with bacterial asparaginase have stimulated efforts to
either subject it to protein engineering or identify other natural
variants. First, L-asparaginase also hydrolyzes L-glutamine
(L-Gln) generating glutamic acid and ammonia, resulting in
neurotoxicity. Second, antibodies generated by patients to
bacterial L-asparaginase decreases the drug’s effectiveness
over time. Development of allergies is thus a significant
hurdle in the clinical use of bacterial L-asparaginase. When
repeated administrations engender a high incidence of allergic
reactions, switching from one bacterial (Escherichia coli) Lasparaginase to another (Erwinia chyrsanthemi) alleviates this
problem somewhat, although the latter is not an ideal substitute for the E. coli enzyme (Duval et al. 2002) possibly due its
shorter half-life (Asselin et al. 1993).
Hyper-sensitivity reactions observed with the bacterial
L-asparaginase might not be as pronounced with fungal
homologs, given the evolutionary relatedness of fungi
and animals as revealed by molecular phylogenetic studies
(Baldauf and Palmer 1993). Based on this premise, we and
others have examined fungi for their L-asparaginase variants.
While Sarquis et al. (2004) reported production of Lasparaginases by Aspergillus, Penicillium and Fusarium, we
found that the enzyme is secreted by fungi endophytic
in marine algae including species of Alternaria, Chaetomium,
Cladosporium, Colletotrichum, Curvularia, Nigrospora,
Paecilomyces, Phaeotrichoconis, Phoma and Pithomyces
(Thirunavukkarasu et al. 2011). Notably, our agar plate-based
colorimetric, qualitative screens revealed that the Lasparaginase variants from Colletotrichum acutatum,
Curvularia lunata, C. eragrostidis, Nigrospora oryzae
and Phomopsis sp. showed selectivity towards L-Asn
over L-Gln (Suryanarayanan, unpublished observations).
If kinetic studies provide a quantitative measure of the
preference for L-Gln over L-Asn, isolating and characterizing these endophytic fungal L-asparaginase variants
would yield a protein-based drug for acute lymphoblastic leukemia with little neurotoxicity. Such efforts
would also complement ongoing protein-engineering
initiatives that seek to increase the asparaginase/glutaminase activity ratio of bacterial L-asparaginase (Offman et al.
2011) and identification of archaeal variants with desirable
attributes (Bansal et al. 2011). Moreover, if the bacterial
version is to be discontinued due to its allergic reactions, the
fungal variants will provide suitable substitutes that could
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become part of a therapeutic regimen in which endophyte
asparginases with different structural epitopes are used
sequentially after the bacterial variant.
Chitin-modifying enzymes
Chitin, the second most abundant biopolymer after cellulose,
is found in the exoskeleton of crustaceans/insects and in
fungal cell walls. Chitin consists of repeating units of β
(1→4)-linked N-acetyl-D-glucosamine and is a waterinsoluble, linear polymer (Fig. 1). Numerous clinical applications for chitin derivatives have evoked a renewed interest in
chitin-modifying enzymes (chitinases, chitin deactylases and
chitosonases). Since fungi produce these enzymes to restructure their chitinous cell wall during growth and (Adams 2004)
and plant infection (El Gueddari et al. 2002), the fungal
enzymes that act on these chitin variants have garnered attention with a view to enlisting these catalysts for reshaping of
naturally occurring chitin to suit specific needs.
The deacetylation of chitin generates a polymer of Nacetyl glucosamine and glucoasamine termed chitosan
(Fig. 1), which is water soluble unlike the parental chitin.
Chitosans are heterogeneous due to variations in their (i)
degree and pattern of N-acetylation (since not all N-acetyl
glucosamine units are converted to glucoasamine), and (ii)
molecular weight. Partially hydrolyzed chitosans are called
chito-ologosaccaharides (CHOS); Aam et al. (2010) suggest
an upper-size boundary for CHOS by restricting the use of
chitin and chitosan to polymers longer than 100 units. Both
chitosans and CHOS are positively charged below pH 6,
with implications for their role as carriers of polyanionic
cargo such as nucleic acids.
While chitin and chitosans have been shown to enhance
the anti-inflammatory response, CHOS have been documented to exhibit antimicrobial, hypo-cholesterolemic,
immuno-stimulating, and anti-tumor/-cancer activities
(Aam et al. 2010; Xia et al. 2011). Chitin derivatives are
highly biocompatible, biodegradable and non-toxic, thus
making them a versatile biomaterial for different biomedical
applications (Table 1). We discuss two examples. First,
some tissue-replacement strategies involve utilization of an
artificial matrix that offers an ideal spatial environment
(porosity, shape and size) for specific stem cells to anchor,
differentiate and divide in the presence of appropriate cues
(Nettles et al. 2002). If such a scaffold also has the added
benefits of antimicrobial and anti-inflammatory properties
(as is the case with chitin, chitosans and CHOS), it is a
bonus for tissue-engineering efforts. Second, CHOS have
been investigated as DNA delivery vehicles. The twin
objectives of such carriers are to stably encapsulate the
desired DNA and deliver the complex to the appropriate
cell/tissue. DNA:CHOS complexes that entail the use of
smaller and narrow size-distributed CHOS were shown to
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Fig. 1 Chitin deacteylasecatalyzed generation of chitosan
from chitin
offer an efficient oral DNA delivery platform since weaker
association (due to fewer positive charges on the smaller
CHOS) promotes efficient release upon delivery without
compromising initial complexation (Köping-Höggård et
al. 2004).
Successful pursuit of the above mentioned applications
require the ready availability of near-homogeneous and
appropriately-sized chitin derivatives, a non-trivial undertaking. Although chemical methods have been employed
to depolymerize chitin/chitosan and deacetylate chitin,
the randomness of these processes contributes to the
heterogeneity of the end products. Enzymatic conversion
of chitin→chitosan→CHOS is preferable due to at least
two reasons–first, we can produce molecules with specific a
number/pattern of N-acetylated residues and a defined molecular weight distribution; second, it is an environment-friendly
approach (Liu and Bao 2009; Aam et al. 2010).
Some filamentous fungi (Aspergillus nidulans, Colletotrichum lindemuthianum, Metarhizum anisopliae, Mucor rouxii,
and Rhizopus nigricans) have been studied for their chitin
deacetylases (Zhao et al. 2010). In our study, a Pestalotiopsis
sp. and a Sordaria sp. that are endophytic in tropical forest
trees were observed to produce high levels of chitin deacetylase (Nagaraju et al. 2009). The chitin deacetylase from Pestalotiopsis sp. acted on colloidal chitin, chitosan with 56 %
degree of acetylation and even a chitin hexamer (Nagaraju et
al. 2009). We recently reported that various fungal endophytes
isolated from forest trees (Alternaria alternata, Aureobasidium pullulans, Botrytis sp., Colletotrichum acutatum, Nigrospora oryzae, Trichoderma sp., and Xylaria spp.) produce
chitosanses which acted on chitosans of different degrees of
acetylation (Govinda Rajulu et al. 2011). Although the characterization is still ongoing, these findings are likely to be
extended to endophytes isolated from marine algae and seagrasses which produce chitin modifying enzymes (Table 3,
Venkatachalam, Thirunavukkarasu and Suryanarayanan,
unpublished).
Since chitin-modifying enzymes have long been studied
from different microbial sources, it is reasonable to inquire
what unanticipated benefits might result from studies of
endophytic fungal strains. Fungal chitosans are better suited
than crustacean chitsoans as scaffolding materials in tissue
engineering since fungal chitosans are of lower molecular
weight, have higher polydispersity and a lower degree of
acetylation (Nwe et al. 2009); the last attribute engenders
various desirable features including smaller pore size, greater mechanical strength and cellular activities (Thein-Han
and Kitiyanant 2006). Consistent with the revised view that
chitosans can be extracted from the cell walls of not only
zygomycetes but from ascomycetes (Hu et al. 2004), we
have shown that ascomycete endophytes produce chitosanases that act on chitosans, whose fraction of acetylated
residues ranges from 38 % to as low as 1.6 % (Govinda Rajulu
et al. 2011). Once a clearer picture emerges of the make-up of
the chitinous cell walls in different endophytes, candidate
organisms with divergent structures could be studied for their
chitin deacetylases and chitosanases that likely contribute to
reshaping the respective chitins/chitosans. It is conceivable
that exploiting the combinatorial capabilities of different
chitin-modifying enzymes from an array of fungal endophytes
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Table 3 Dot blot-based identification of chitin-modifying activities in fungal endophytes
isolated from seagrasses and
seaweeds
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Substrate used
Number of fungal endophyte isolates
tested
Positive for activity (%)
Glycol chitin
102
14 %
Chitosan; 1.6 %, degree of acetylation
102
39 %
Chitosan; 38 %, degree of acetylation
70
66 %
Chitosan; 56 %, degree of acetylation
102
57 %
will help generate ‘designer chitosans’ needed for specific
applications (Nagaraju et al. 2009).
Utility of endophytic biotransformations in understanding
drug metabolism and discovery of new drugs
Fungi perform several biotransformation reactions including
stereo-selective hydroxylation, sulfoxidation, expoxidation,
and deracemization (Borges et al. 2009). Due to their ability
to detoxify the defense chemicals of their host plants, it is
perhaps not surprising that endophytes catalyze biotransformation of a variety of molecules (Shibuya et al. 2005; Verza et
al. 2009; Borges et al. 2009; Barth et al. 2010; Zikmundova et
al. 2002). For instance, endophytes isolated from Aphelandra
tetragona biotransfomed phytoanticipin molecules to novel
metabolites through acylation, oxidation, reduction, hydrolysis, and nitration reactions (Zikmundova et al. 2002). Of
particular relevance is the fact that fungal endophytes are
capable of transforming drugs to products similar to those
formed by phase I metabolism of xenobiotics in the mammalian liver. An important payoff then relates to their ability to
generate putative mammalian metabolites in large quantities
for subsequent isolation and pharmoclogical/toxicological
testing in animal models. For example, endophytic isolates
of Phomopsis, Glomerella, Diaporthe and Aspergillus transform with high regio- and stereoselectivity thioridazine, a
neuroleptic drug, to subsets of products produced by mammalian metabolism (Borges et al. 2008). Similarly, Pestalotiopsis guepini transformed norfloxacin, a fluoroquinolone
anitmicrobial, to metabolites that are produced by mammalian
cells (Parshikov et al. 2001).
Since these fungal endophyte-catalyzed biotransformations
are enantio-selective, the production of chiral compounds
from racemic mixtures is an appealing feature (Borges et al.
2007). Biotransformations also play a major role in drug
discovery by modifying lead molecules to less toxic variants
without compromising biological activity of the lead (Shu et
al. 2008). This expectation is exemplified in the work of Tang
et al. (2011) who used the endophytic Alternaria alternata and
plant pathogenic Gibberella fujikuroi to perform a tandem
biotransformation of podophyllotoxin (a lignan with anticancer activity but exhibiting high toxicity to humans) to a novel
derivative, 4-(2,3,5,6-tetra-methylpyrazine-1)-4-demethyl-
epipodophyllotoxin, which displayed high antitumor activity
but low cytotoxicity.
Many virulent plant pathogenic fungi have the ability to
detoxify different types of phytoalexins, which are defense
metabolites produced by host plants in response to biotic
stresses including a fungal infection (Pedras et al. 2005).
Different fungi can produce distinct enzymes to detoxify the
same phytoalexin; for example, Alternaria brassicicola,
Sclerotinia sclerotiorum and Leptosphaeria maculans produce brassinin hydrolase, brassinin oxidase and brassinin
glucosyl transferase, respectively, for detoxifying the phytoalexin brassinin of crucifers (Pedras and Minic 2011).
Since some endophytes are also latent pathogens (Photita
et al. 2004; Suryanarayanan and Murali 2006), they are
likely to express a repertoire of plant metabolite-detoxifying
enzymes, which could be drafted for biotransformations.
Endophyte enzymes in food and nutrition
Since many microbial enzymes have been exploited for applications in the food industry, why should variants isolated from
fungal endophytes deserve special attention? Wide-ranging
industrial applications necessitate use of enzymes with different properties thus lending significance to isolation of catalysts from diverse niches. In this regard, we point out that there
is an increasing appreciation that fungal endophytes do exhibit
traits that would be expected of their plant host growing in a
given habitat. For example, endophytes isolated from plant
species in the Baima Snow Mountain (Li et al. 2012) exhibit
cold adaptation suggesting that these fungi might harbor coldattuned enzymes well suited for specific applications including biotransformation of heat-labile compounds that can only
be performed at low temperatures. In a related vein, a salttolerant protease (of interest to the fish/soy sauce manufacturing process) was purified from Aspergillus oryzae LK-101,
which had been isolated from soybean paste (Lee et al. 2010).
Our own findings on endophytic fungal tannases also confirm
these observations.
Tannase (also called tannin acylhydrolase) is induced by
tannic acid in some fungi such as Aspergillus and Penicillium (Ramírez-Coronel et al. 2003) and is used as a clarifying
agent in the manufacture of instant tea, wine and fruit juices.
Tannases hydrolyze tannic acid to release gallic acid, glucose
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and galloyl esters of glucose. Since tannins are generally antifungal in nature (Dix 1979), it is reasonable to expect endophytes residing in tannin-rich tissues to have evolved mechanisms to tolerate tannins. We discovered that fungal
endophytes such as Colletotrichum sp., Paecilomyces sp.,
Phoma sp., Phomopsis sp. and Phyllosticta sp., isolated from
tannin-rich mangrove leaves were able to grow on tannic acidamended medium (Kumaresan et al. 2002). Moreover, we
observed that endophytes residing in tannin-rich leaves of
Manigifera indica grew better on medium containing tannic
acid than conspecific endophytes from leaves of Zizyphus
xylopyrus, which have less tannin (Mohan Doss and Suryanarayanan unpublished). This finding illustrates the utility of
isolating endophyte tannanases specifically from the plant
whose tannin needs to be biotransformed. Parenthetically,
we note that fungal endophytes can use tannins as a sole
carbon source (data not shown); this is not unanticipated given
that fungi can degrade tannins to aliphatic amino acids, which
are then catabolized to intermediates in the Krebs cycle (Watanabe 1965; William et al. 1986; Bhat et al. 1998).
While the above examples clearly illustrate that enzymes
from fungal endophytes might offer benefits for hydrolyzing
certain substrates under defined conditions, it is important to
first build an inventory of industrially-relevant enzymes
produced by tropical fungal endophytes (e.g., proteases,
lipases and amylases). In this regard, our preliminary findings are encouraging in demonstrating the presence of
enzymes of interest to the food industry in a wide range of
fungal endophytes. However, further purification and characterization of these variants is critical before adapting any
of them for a specific application.
While bacterial alkaline protease is extensively used in
feed, denture cleansers, baking, and brewing (Gupta et al.
2002), acid proteases are used in the production of seasoning materials, fermented food, and digestive aids (Rao et al.
1998). The most common source of acid protease is Aspergillus oryzae; attempts are ongoing to enhance its yield and
activity by optimizing culture conditions and subjecting it to
mutagenesis (Hideyuki et al. 2002). In our studies using
agar plates, we found that 59 % of the 134 foliar endophytes
isolated from dicotyledonous trees were positive for protease activity (Table 2). A dot-blot assay to identify the type of
proteases (acidic, neutral or alkaline) revealed that several
endophytes including species of Colletotrichum, Corynespora, Curvularia, Nodulisporium, Robillarda and xylariaceous fungi produced alkaline proteases; many of these were
also positive for acidic proteases (Thirunavukkarasu and
Suryanarayanan, unpublished).
Fungal α-amylases are preferred in bread manufacture over
other sources of this enzyme as they do not discolor the bread;
they are also used extensively in beverage and soft drink manufacture (Arora et al. 2004). Fungal lipases are used in baking,
manufacture of beverages, cheese, butter emulsification, health
Fungal Diversity (2012) 54:19–30
food and fat removal from meat and in the preparation of
digestive aids (Arora et al. 2004). Several endophytic fungi that
we screened produce α-amylases and lipases (Table 2). Amylase
from Aspergillus oryzae is extensively used in baking industry
but it has been reported to induce a respiratory allergy response
in bakery workers (Houba et al. 1996). Akin to our contention
with asparginases of endophytic origin, we suggest that the
endophyte lipases and amylases be studied for potential higher
human compatibility.
Energy and environment
Biofuel production
Escalating energy consumption, rapid depletion of fossil
fuels and environmental concerns have resulted in the search
for alternative next-generation transportation fuels and
renewed interest in plant-derived fuels. The latter entails
(i) the deconstruction of polysaccharides in plant biomass
to constituent sugars, and (ii) the microbial fermentation of
these sugars to alcohols (e.g., ethanol, butanol). To avoid the
adverse effects of diverting food/feed crops for biofuel
production, it is imperative to use as feed stocks noncompeting bioresources, preferably those which utilize nonagricultural land–ideal candidates would be the lignocellulosic
biomass of herbaceous/woody plants and agricultural waste
such as straw, sugarcane bagasse and corncob (Camassola and
Dillon 2009). In this context, we focus on a major technical
roadblock related to the degradation of the cellulosic fraction
of lignocellulosic biomass to fermentable sugars.
Lignocellulose is comprised of cellulose, hemicellulose,
and lignin (variability in the relative proportions depend on
the plant species). While cellulose is a polymer of β(1→4)linked glucose, hemicellulose is a heterogeneous polymer
largely comprised of hexoses (e.g., mannose, galactose,
glucose) and pentoses (e.g., arabinose, xylose). Natural degradation of cellulose/hemicellulose is carried out by the
synergistic attack of various glycohydrolases; endoglucanases that hydrolyze internal glycosidic bonds in cellulose
to form cello-oligosaccharides, which are then exolytically
attacked by cellobiohydrolases to release the cellobiose
disaccharide, which in turn is hydrolyzed to monomeric
glucose units by β-glucosidases. For cellulases to break
down cellulose, they must gain access to the insoluble
cellulose fibrils enmeshed in hemicellulose and lignin. To
overcome this problem, the physical and chemical pretreatment, which precede enzymatic hydrolysis, often make
it difficult to rein in both the cost and the attendant waste
management issues (Mosier et al. 2005).
There has been intense research over the past several
decades with major advances in the commercial use of
cellulases and xylanases (in particular from the fungus Trichoderma reesei) to deconstruct cellulose. However, large-
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Fungal Diversity (2012) 54:19–30
scale industrial adaptation of the process has been thwarted
both by the intrinsic recalcitrance of plant biomass (stemming in part from the inherent variation in cell wall composition) and the limited number of organisms from which the
cellulolytic enzymes have been derived. It has been argued
that efficient and economical saccharification of biomass
requires bioprospecting microbes of less-studied environments such as termite guts, rumen and tropical forests
(Somerville 2007). We rationalize below why fungal endophytes merit inclusion in this list.
Our understanding of plant cell wall-degrading enzymes of
fungi is far from complete. For instance, it is not clear how
Trichoderma reesei, the most efficient producer of industrial
cellulases and hemi-cellulases, has fewer genes encoding
cellulolytic enzymes compared to related Ascomycetes
(Martinez et al. 2008). In comparison, some plant pathogenic
fungi have more cellulases, hemicellulases, pectinases,
carbohydrate-binding modules, carbohydrate esterases, and
polysaccharide lyases; notably, they also produce unique
glycosyl hydrolases that are not coded by T. reesei (Paper et al.
2007; Martinez et al. 2008). It is now appreciated that
enzymes of mutant strains of T. reesei, selected or engineered
for industrial generation of biofuels, may not offer the best
route for degrading cell walls of different plant species owing
to differences in cell wall chemistry (King et al. 2011;
Shrestha et al. 2011). Given this scenario, endophytes
which, akin to plant pathogenic fungi, have to degrade
the plant cell wall to infect and also to compete with a
multitude of litter-degrading organisms (when continuing
their life as a litter fungus) are likely to express a wide
repertoire of glycosyl hydrolases and cellulases, and offer
new catalysts for use in combination with T. reesei–derived
enzyme cocktails. Indeed, a few studies have already shown
that endophytes elaborate efficient plant cell wall-degrading
enzymes (Weber et al. 2004). We also observed that many of
the endophytes isolated from tropical trees produce cellulases
(Table 2; Suryanarayanan, unpublished).
Cellulases of different endophytes warrant investigation
and characterization to explore the possibility of using a
consortium of biocatalysts from different species to facilitate
a more efficient and cost-effective biomass conversion; it is
conceivable that in addition to differences in catalytic efficiency and combinatorial capabilities in depolymerizing
cellulose and hemicellulose, this consortia approach might
benefit from different, possibly complementary, mechanistic
strategies used by these enzymes to loosen the cellulose
fibril structure and enhance substrate molecular disorder, a
pre-requisite for efficient and rapid degradation. Moreover,
since the renewable biomass feed stocks are likely to be
different depending on the geographical location, it would
be prudent to identify the enzymes best suited for the specific
plant being considered for bioenergy. We contend that
enzymes from endophytes isolated from the target plant will
25
likely offer the best deconstruction strategy. Support for
this premise comes from a recent study which demonstrated
elegantly that fungi isolated and cultured from Miscanthus
were indeed able to expedite breakdown of whole alkalitreated and ground Miscanthus (Shrestha et al. 2011).
Even as advances are being made toward the objective of
designing a consolidated bioprocessing and fermentation
platform in which a single microbe can efficiently convert
plant-derived pentoses/hexoses to biofuels, an important
roadblock relates to the presence of fermentation-inhibitory
compounds in lignocellulosic biomass hydrolysates. High
temperature and dilute acid hydrolysis-based pre-treatment
of plant biomass is commonly employed to hydrolyze the
hemicellulose fraction and disrupt the lignin sheath, whose
presence on the surface of cellulose prevents efficient enzymatic hydrolysis of cellulose to glucose. This pre-treatment,
albeit favored for its adaptability to large-scale operations,
unfortunately generates unwanted products such as furfural
and hydroxymethyl furfural (HMF), which are microbial
fermentation inhibitors (Jönsson et al. 1998). Depending
on the biomass source and hydrolysis procedure employed,
these furanic aldehydes can be present at concentrations as
high as 0.1 g/g of lignocellulosic hydrolysates (Almeida et al.
2007). Since microbial growth rates and alcohol production
are adversely affected by these furans, eliminating them from
the fermentation medium is critical to ensure high yields.
While detoxification of these aldehydes is a useful
bio-abatement strategy, of greater utility is the ability to
convert these furans into substances that will contribute
to energy metabolism, with associated payoffs in growth
and fermentation. Such a goal is based on the ability to
identify microbial genes necessary for metabolizing these
furans, and then introducing these furanic aldehyde catabolic
pathway genes for heterologous expression in a microbe already engineered for economical production of biofuels. Motivated by this long-term goal, we recently investigated if
these furans can support the growth of endophytes; although
there are some examples of fungi metabolizing fufural or
HMF (e.g., Coniochaeta ligniaria; López et al. 2004), studies
to date have mostly been restricted only to those that are now
commercially used for cellulose breakdown (Liu 2011).
Our preliminary experiments showed that some of the
endophytes such as Arthrinium sp., Colletotrichum sp., Pestalotiopsis spp. and Sordaria sp., could grow on media with
either furfural or HMF as the sole carbon source
(Suryanarayanan, unpublished). In all these cases, we
observed an initial slower growth rate in the presence
of these aldehydes when compared to growth on sucrose
alone–this difference, however, was diminished significantly after 10 days as evidenced by a near-identical
final diameter of the colony. Interestingly, our findings
on the growth lag and the preference for HMF compared to furfural are reminiscent of earlier reports with
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26
other fungi such as Fusarium oxysporum (Modig et al.
2002; Xiros et al. 2010).
It is vital to perform a detailed characterization of the
catabolic pathways/enzymes used by endophytes to convert
furfural/HMF to an energy-yielding intermediary metabolite. Although availability of the target endophyte genome
sequence per se will not address this question, comparison
with established routes e.g., the furoic acid-centered Trudgill pathway in Gram-negative aerobic bacteria that converts
furfural to α-ketoglutarate (Trudgill 1969; Koopman et al.
2010) might reveal parallels and differences. As pointed out
by Wierckx et al. (2011), bioabatement processes based on
catabolic pathways from aerobic bacteria such as Cupraividus basilenesis necessitate aeration and near-neutral pH,
with the former increasing cost and latter the risk of infection. Given the ability of endophytic fungi to function as
litter degraders, we are also investigating the ability of
endophytes to grow on furfural and HMF at acidic pH.
Another facet to biofuel production is the possibility of
using brown seaweeds/macroalgae as a feedstock, one
whose appeal is enhanced by the fact that it does not require
land, fresh water or fertilizer. In addition to mannitol and
glucans, these seaweeds have as a structural polymer alginate, a polysaccharide made of guluronic and mannuronic
acids. Since alginates cannot be converted to ethanol by the
microbes, Wargacki et al. (2012) successfully engineered E.
coli (by reconstructing a pathway from Vibrio splendidus) to
degrade, transport and metabolize alginates thus paving the
route for direct biofuel production from seaweeds. Since we
have observed that many fungal endophytes (e.g., Aspergillus
niger, A. terreus, Penicillium sp.) are associated with brown
seaweeds (Suryanarayanan et al. 2010) and since alginate
lyase has been reported from a fungus associated with the
brown seaweed (Singh et al. 2011), screening seaweedassociated endophytes for novel alginate utilization pathways
might prove valuable to the biofuel industry.
Bioremediation
Chitin and its modified products have been used to detoxify
waste water (Bhatnagar and Sillanpää 2009). For example,
modified and chemically crosslinked chitosans selectively
adsorb heavy metal ions from aqueous solutions (Kandile
and Nasr 2009), and chitosan resins absorb uranium (VI)
(Zhou et al. 2012). As stated earlier, the diversity of chitinmodifying enzymes in fungal endophytes should be
exploited to generate chitosans of desired structures, which
would serve as environment-friendly bio-absorbents that
can help pollution control.
Fungal laccases have wide substrate specificity and act
on many small organic substrates including polyphenols,
methoxy-substituted phenols, and aromatic amines. They are
used in paper manufacture for delignification, bioremediation
Fungal Diversity (2012) 54:19–30
of phenolic compounds and biobleaching (Kunamneni et al.
2008), and in pretreatment of lignocelluloses for biofuel production (Piscitelli et al. 2011). The commercial use of laccases
is limited due to its low expression levels and catalytic efficiency (Kunamneni et al. 2008). Fungal endophytes are
known to produce laccases (Kumaresan et al. 2002;
Promputtha et al. 2010). We observed that more than 60 %
of the 134 endophyte isolates tested were positive for laccase.
The white rot basidiomycete fungi secrete highly efficient
laccases when compared to the ascomycete fungi (Rodgers
et al. 2010). Although most of the endophytes belong to the
ascomycetes, endophyte laccases remain to be characterized
before such a generalization can be made. Such a contention is
supported by the case of lignolytic enzymes, which are commonly used in bio-bleaching and degradation of recalcitrant
organ pollutants. Although it was widely held that the white
rot fungi are distinctive for their lignolytic enzymes,
Urairuj et al. (2003) showed that several xylariaceous
endophytes (ascomycetes) host such enzymes. Thus, a
systematic survey of endophytes, especially those that
switch to a saprobic phase as litter degraders in senescent
tissues, for the presence of such laccases and ligninases, could
furnish new insights into fungal metabolism and contribute to
improved bioremediation methods.
Recent reports have revealed some unexpected bioremediation opportunities based on fungal endophytes. We illustrate with two examples. First, plastic waste reduction, a
shared global objective, could be aided by the identification
of microbes that catabolize polyurethane. Indeed, Russell et
al. (2011) showed that a Pestalotiopsis isolate produces a
polyurethanase (member of the serine hydrolase family),
which can degrade polyester polyurethane, and help use this
polymer as the sole carbon source under aerobic and anaerobic
conditions. Second, high-temperature pyrolytic industrial processes are known to contribute to the high environmental
levels of polycyclic aromatic hydrocarbons (PAHs); given
the carcinogenic properties of PAHs and risk to human
health, decreasing PAHs levels in the environment is
highly desirable. Remarkably, Dai et al. (2010) demonstrated that endophytic Ceratobasidum stevensii isolated
from Bischofia polycarpam could degrade phenanthrene, a
PAH, thus highlighting a new bioremediation application for
fungal endophytes.
Challenges and prospects
The co-evolution of fungal symbionts and their plant hosts,
during an association that dates back to at least the preCretaceous period (Krings et al. 2012), is likely the underlying basis for the potent and diverse arsenal of enzymes in
fungal endophytes (De Fine Licht et al. 2010). We suggest
an inter-related two-pronged approach to exploit this rich
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Fungal Diversity (2012) 54:19–30
collection of enzymes for new applications and to further
our understanding of biocatalysis.
First, while there are ongoing efforts to improve the performance of currently used fungal enzymes by methodologies
including site-directed mutagenesis, directed evolution, and
semi-rational design/protein engineering (Heckmann-Pohl et
al. 2006; Piscitelli et al. 2011), there is a need to undertake
extensive bioprospecting to identify fungi producing more
efficient and novel industrial enzymes. In this context, we
stress that identification of candidate organisms from a huge
assemblage of endophytes from any given ecological niche
will be a daunting task. A plant host-guided search for endophytes that produce enzymes of desirable physical, chemical
and biological traits could be beneficial. Endophytes in aerial
and root tissues of plants growing in extreme environments
(experiencing extreme pH, temperature, high salt or metal
concentrations) may secrete enzymes adapted to such harsh
environments, which unlike many of their mesophilic counterparts might find use in specific industrial processes. We consider a few examples here. We recently discovered that spores
of litter fungi from a tropical forest (which also survive as
endophytes in the trees of this forest) that experiences prolonged drought and seasonal fires are extremely heat-resistant
and can survive sudden exposure to dry heat of over 100 °C
for 2 h (Suryanarayanan et al. 2011). Although their mycelia
are mesophilic, it would be worth examining heat-treated
spores of these endophytes for the presence of thermostable
enzymes. Similarly, psychrophilic endophytes from highaltitude plants (Li et al. 2012) are likely to be good source of
cold-adapted enzymes. Also, salt-tolerant fungal endophytes
of mangroves (Kumaresan et al. 2002) and marine algae,
which synthesize certain metabolites only in the presence of
salt (Suryanarayanan 2012), could be good candidates for
identifying salt-tolerant enzymes. Although similar expectations of archaea, which colonize extreme environments, have
not translated into many biotechnological applications, a key
distinction is the ease with which endophytes can be cultured
in a large scale outside the plant. From crude extracts of
endophytes, specific enzymes of interest under desired assay
conditions could be identified using either a rapid, quantitative
and high-throughput screen (King et al. 2009) or even qualitative tests such as dot blots (e.g. Govinda Rajulu et al. 2011),
filter centrifugation (Heinonsalo et al. 2012) and agar plate
assays (Kumaresan et al. 2002). As stated earlier, these exercises become even more meaningful if the industrial application involves a certain plant substrate (e.g., tannins), since the
search for an appropriate enzyme could focus on the endophytes associated with this plant.
Second, genome mining could prove valuable in discovering novel enzymes from endophytes (Kaplan et al. 2011).
Increasing the number of genome sequences and thus the
amino acid sequences (and even predicted tertiary structures) will help us to better appreciate and further
27
manipulate the unique physicochemical or biological characteristics of enzymes from fungal endophytes. Despite the
rapid decrease in costs associated with the development of
third- and fourth-generation sequencing methods, it would
be practical to first focus on a few organisms to be sequenced. While this choice might be dictated in part by
the first approach (described above) that identifies one or
two endophytes producing the most number of enzymes of
commercial interest, we provide an alternative rationale that
might expediently achieve this goal. The endophyte-plant
association has existed for millions of years resulting in the
evolution of multi-host endophytes (Krings et al. 2012).
Endophyte genera such as Phomopsis and Pestalotiopsis
occur in phylogenetically-distant and geographicallyseparated host plants (Suryanarayanan 2011). We need to
understand what attributes contribute to the ecological success and broad host range of these fungi. It is worth emphasizing that most endophytic Phomopsis isolates produce
enzymes including cellulases, lipases, pectinases, pectate
lyases and proteases. We have also observed that species
of Phomopsis endophytes produce β-glucosidases (Firkins,
Gopalan and Suryanarayanan, unpublished), tannases
(Mohan Doss and Suryanarayanan, unpublished) and Lasparaginases lacking glutaminase activity (Suryanarayanan
et al. unpublished). Furthermore, as mentioned earlier,
Phomopsis sp. could also utilize HMF as the sole carbon
source. Similarly, Pestalotiopsis endophytes elaborate novel
chitin deacetylases (Nagaraju et al. 2009), have conidia that
are highly thermotolerant (Suryanarayanan et al. 2011), and
produce a variety of secondary metabolites including alkaloids, terpenoids, isocoumarin derivatives, coumarins, chromones, quinones, semiquinones, peptides, xanthones,
xanthone derivatives, phenols, phenolic acids, and lactones
(Xu et al. 2010). It is conceivable that host range expansion
of these cosmopolitan endophytes (Phomopsis sp.,
Pestalotiopsis sp.) is due to their ability to produce a
broad range of enzymes in response to environmental
cues. Therefore, these endophytes must be given a high
priority for genome sequencing if we are to exploit their
remarkable catalytic versatility. Since the currently existing fungal genome sequences include only those of
mycorrhizae, plant pathogens and some industrially important fungi (Porras-Alfaro and Bayman 2011), there is
also considerable immediacy to sequencing endophytes
from the perspective of having genomes from every
branch of the tree of life.
Acknowledgments We are grateful to Drs. E. J. Behrman and T.
Ezeji (OSU) for comments on the manuscript, and to Andrew Wallace
(OSU) for preparing Fig. 1. TSS thanks the United States-India Educational Foundation (USIEF), New Delhi and the Fulbright Scholar
Program (USA) for the award of a Fulbright-Nehru Senior Researcher
grant to characterize fungal endophyte enzymes in VG’s laboratory at
OSU, and the Department of Biotechnology, Government of India for
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28
funding the Indo-German Research Project BT/IN/FRG/09/TSS/2007
on endophyte enzymes. VG gratefully acknowledges funding support
from the Northeast Sun Grant Initiative Award 52110–9615 from US
Department of Transportation (via a sub-contract from Cornell University
to T. Ezeji and V. Gopalan, OSU).
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