RNA-sequencing reveals the complexities of the transcriptional response to lignocellulosic biofuel substrates in Aspergillus niger.

BACKGROUND: Saprobic fungi are the predominant industrial sources of Carbohydrate Active enZymes (CAZymes) used for the saccharification of lignocellulose during the production of second generation biofuels. The production of more effective enzyme cocktails is a key objective for efficient biofuel production. To achieve this objective, it is crucial to understand the response of fungi to lignocellulose substrates. Our previous study used RNA-seq to identify the genes induced in Aspergillus niger in response to wheat straw, a biofuel feedstock, and showed that the range of genes induced was greater than previously seen with simple inducers.
RESULTS: In this work we used RNA-seq to identify the genes induced in A. niger in response to short rotation coppice willow and compared this with the response to wheat straw from our previous study, at the same time-point. The response to willow showed a large increase in expression of genes encoding CAZymes. Genes encoding the major activities required to saccharify lignocellulose were induced on willow such as endoglucanases, cellobiohydrolases and xylanases. The transcriptome response to willow had many similarities with the response to straw with some significant differences in the expression levels of individual genes which are discussed in relation to differences in substrate composition or other factors. Differences in transcript levels include higher levels on wheat straw from genes encoding enzymes classified as members of GH62 (an arabinofuranosidase) and CE1 (a feruloyl esterase) CAZy families whereas two genes encoding endoglucanases classified as members of the GH5 family had higher transcript levels when exposed to willow. There were changes in the cocktail of enzymes secreted by A. niger when cultured with willow or straw. Assays for particular enzymes as well as saccharification assays were used to compare the enzyme activities of the cocktails. Wheat straw induced an enzyme cocktail that saccharified wheat straw to a greater extent than willow. Genes not encoding CAZymes were also induced on willow such as hydrophobins as well as genes of unknown function. Several genes were identified as promising targets for future study.
CONCLUSIONS: By comparing this first study of the global transcriptional response of a fungus to willow with the response to straw, we have shown that the inducing lignocellulosic substrate has a marked effect upon the range of transcripts and enzymes expressed by A. niger. The use by industry of complex substrates such as wheat straw or willow could benefit efficient biofuel production.

Background

Low molecular weight organic acids production by filamentous fungi, have attracted considerable attention for their role in natural ecology and their potential industrial applications [1],[2]. Fungal natural production of organic acids is thought to have many key roles in nature depending on the type of fungi producing them. These roles are either due to the pH decrease consecutive to their secretion or to direct interaction of the organic acid with the environment [3],[4]. The consecutive decrease in pH upon their secretion may give a competitive advantage to the acid-tolerant filamentous fungi. For ectomycorrhizal fungi, this pH decrease also has been suggested to solubilize soil minerals thus releasing nutrient ions for uptake by plants and microorganisms, enhancing mineral weathering [1]. For saprophytic and wood-decaying fungi, this pH acidification, caused by oxalic acid production, leads to an acid-catalyzed hydrolysis of holocellulose [5]–[7]. Concerning their direct interaction with the environment, organic acids participate in metal detoxification by metal complexion and oxalic acid plays a major role in biomass degradation [4]. For this reason, Basidiomycota have been extensively studied for their ability to produce oxalic acid [8]–[12]. To better understand their role in the ecosystem, these studies have focused on plant/fungi symbiosis [9],[13], or growth on complex substrates [12],[14]–[16], and are often focused on wood-decay or mycorrhizal fungal species.

In addressing the demand for sustainable alternatives to fossil fuel as a source of energy and chemicals, synthetic biology focuses on understanding how biological systems work and how to use them to benefit society. Organic acids can have multiple industrial applications as food additives, pharmaceutical and cosmetic excipients [17]. They are fully degradable molecules and can be used as chemical intermediates or as synthons for the production of biodegradable polymers, potentially replacing petroleum-based or synthetic chemicals [17]. Some fungi are well known for their natural capability to produce high amounts of various useful organic acids. These fungi are mostly from the Aspergillus (e.g. citric, gluconic, malic and itaconic acids) and Rhizopus genera (e.g. lactic and fumaric acids). Some of these organic acids (i.e. citric acid) can be produced through large-scale bioprocesses, showing the high potential of fungi as organic acid production plateforms [2],[18].

The literature concerning organic acid production in filamentous fungi often focuses on one specific organic acid and there is little information about the other metabolite produced. In many cases these studies focus on specific strains and cultures are carried out in different conditions and with different complex media. Therefore, it is sometimes difficult to compare the potentiality of filamentous fungi from the literature. Moreover, the fungal biodiversity is estimated to be 1.5 million species [19] and there is still a lot to learn about their potential for metabolite production. In this study, 66 strains of saprophytic and wood-decay fungi (40 Ascomycota and 26 Basidiomycota) were selected and studied in liquid glucose medium, without pH regulation, in order to compare their metabolic features. These strains belong to 47 different species, representing 23 fungal families. The majority of the strains tested were collected in situ from different geographical areas such as tropical forests from French overseas territories and temperate forests from metropolitan France [19]. Fungal growth and metabolites production was done in glucose liquid media without any pH regulation to take into account industrial up- and down-stream technical and economical issues. These conditions, close to industrial ones, were chosen to highlight the potential of these organisms for industrial organic acid production. The great diversity and origin of the selected strains enable us to compare the potentiality of a number of fungal groups for the production of organic acids and ethanol.

Results and discussion

Growth of the selected strains and pH of the medium

All the Ascomycota were able to grow in the liquid medium at an initial pH of 5.5. However, 4 strains of Basidiomycota: Ischoderma benzoinum (BRFM1133), Grifola frondosa (BRFM1162), Panellus serotinus (BRFM1284), and Polyporus squamosus (BRFM1531), did not grow to a sufficient level and did not acidify the medium. These strains were not considered for the following steps. The pH of the medium was acidified for most of the cultures but to different extents. An extreme acidification, to pH below or equal to 2, was observed for 33 Ascomycota strains, representing 80% of the strains from this phylum. In particular, all the Aspergilli tested (22 strains) acidified the medium below pH 2 (Figure 1). Five strains of Ascomycota acidified the medium to pH between 2 and 4 and only two strains did not or slightly acidify the medium. To the contrary, only two Basidiomycota highly acidified the medium to pH below or equal to 2, namely Phanerochaete chrysosporium (BRFM413) and Trametes menziesii (BRFM1281). Eight strains acidified the medium between 2 and 4, and 12 strains did not or slightly acidified the medium (Figure 1). For the strains acidifying the medium, the acidification started within 24 hours of growth and the final pH was already observed after 3 days of growth.

Figure 1

Repartition of and strains according to the final pH of growth medium after 6 days of incubation. (orange square) Ascomycota, (sky blue square) Basidiomycota.

Organic acid and ethanol production

HPLC analysis of the supernatants obtained at day 3 of incubation contained only oxalic, malic, propionic and citric acids, found mostly in the supernatants of Aspergillus species. The samples taken at day 6 of incubation showed a better view of the potentiality of the strains tested for organic acid production and allowed the detection of 15 different carboxylic acids at concentration between 0.1 and 3.7 g.L−1: acetic, ascorbic, butyric, citric, fumaric, formic, oxalic, gluconic, itaconic, isobutyric, lactic, malic, propionic, succinic, and tartaric acids (Figure 2). Ethanol was the main neutral metabolite.

Figure 2

Hierarchical clustering of organic acids and/or ethanol producing strains. Concentration were determined by HPLC-UV or RI analysis and expressed as a percentage of the maximum concentration observed for each metabolite and represented by a color scale with different intensity of blue. Concentration of butyric, tartaric, oxalic, malic, citric, gluconic, succinic acids and ethanol were used to build distance tree. The figure was edited using the Multiexperiment Viewer software [20].

Hierarchical clustering was used to classify the strains producing detectable amounts of organic acids and/or ethanol at day 6 of incubation. The clustering was based on production levels of the most widely detected compounds: butyric, citric, gluconic, malic, oxalic, succinic, tartaric acids and ethanol (Figure 2). Two main groups appeared in this clustering: one group of organic acid and ethanol producers and one group producing mainly ethanol.

In the first group, which was mainly composed of Aspergilli, all the compounds analyzed in our assays were detected. This first group represented fungi co-producing a wide variety of organic acids at relatively high concentrations. Two sub-groups could be observed. The first one was composed of various Aspergilli and Nectria species, with only three A. niger species. The second one was composed mostly of A. niger species, with the exception of A. terreus (BRFM111). These results show that A. niger is clearly an exception in the fungal kingdom concerning organic acid production. Organic acid production may also be used along with secondary metabolites in the chemotaxonomy of Aspergilli [21].

The second group was composed of the remaining Ascomycota and Basidiomycota producing mainly ethanol. The metabolite concentrations obtained were lower than for the first group and the variety was narrower since only 6 different organic acids out of 15 were detected (Figure 2). In this second group, two subgroups were observed with strains producing only ethanol and strains producing ethanol and/or other organic acids. Ethanol was detected in 33 strains out of 40 regardless their phylum and species. Although this ethanol production is surprising for organisms traditionally considered as non-fermentative, there are previous records of ethanol production by filamentous fungi. Recent literature shows an increasing interest in ethanol production by filamentous fungi, in particular Flammulina velutipes [22],[23]. Some species belonging to Fusarium, Mucor and Paecilomyces were also found to efficiently convert xylose to ethanol with high yields [24]-[26].

Concerning the relation between pH acidification (section Growth of the selected strains and pH of the medium) and organic acid production, as expected most of the highly acidifying strains were good organic acid producers, from the Ascomycota phylum. However, the pH obtained in the Ascomycota growth media was below 2, which is far below the pKa of organic acids (between 3 and 5). Moreover, some strains for example Phanerochaete chrysosporium (BRFM413) and Cosmospora vilior (BRFM1198) acidified the medium below 3 but did not produce detectable amounts of organic acids. Therefore, the decrease in pH in our experiments cannot be explained by the sole release of large amount of organic acids. The acidification of the medium is probably mainly due to the removal of the ammonium from ammonium sulfate salt, used as nitrogen source, or excretion of H+ ions from the assimilation of NH4 +.

Organic acid and ethanol production in the Basidiomycota phylum

In the Basidiomycota phylum, only 6 strains out of 20 produced organic acids or ethanol. Pycnoporus coccineus (BRFM1396) was the only Basidiomycota producing several metabolites with 0.6 g.L−1 of gluconic acid, 0.2 g.L−1 of formic acid and 0.2 g.L−1 of ethanol (Figure 2). Stereum hirsutum (BRFM889), Tinctoporellus epimiltinus (BRFM1229) and Fomitiporia mediterranea (BRFM1315) produced only ethanol at concentrations between 0.12 and 0.19 g.L−1. Two strains: Postia stiptica (brown-rot, BRFM1148) and Ganoderma weberianum (white-rot, BRFM1548) produced only oxalic acid at 0.06 and 0.08 g.L−1, respectively. Generally, oxalic acid is accumulated in large quantities by brown-rot fungi and detected in lower amounts in white-rot fungi [8],[10]. This difference was attributed to the inability of brown-rot fungi to undertake an active regulation of oxalic acid concentration [10]. The other brown-rot tested, Gloeophyllum sepiarium, produced neither organic acids nor ethanol. Interestingly, P. coccineus (white-rot) produced 0.2 g.L−1 of formic acid. Formic acid production by this strain might be the result of oxalate decarboxylation, as described previously for white-rots [11].

Organic acid and ethanol production in the Ascomycota phylum

Out of the 40 strains, 6 strains of Ascomycota produced neither organic acids nor ethanol at detectable level: Cosmospora vilior (BRFM982 and BRFM1198), Nectria pseudocinnabarina (BRFM1288), Xylaria schweinitzii (BRFM1447), Hypomyces luteovirens (BRFM1580), and Cordyceps militaris (BRFM1581). Ascorbic, fumaric and itaconic acids were detected in only a few supernatants, all from Aspergilli, and at concentrations below the limit of quantification (0.05 g.L−1). The maximal metabolite concentrations were all observed in Aspergilli culture media and ranged from 0.1 g.L−1 for lactic and tartaric acids to more than 2 g.L−1 for citric, formic, gluconic acids and ethanol (Table 1). Indeed, among the fungal kingdom, Aspergilli are well known for their ability to accumulate large amounts of organic acids [2],[18]. In our culture conditions the concentration obtained were low compared to the literature where the conversion of glucose into organic acid is described to approach 100% for some Aspergilli in optimized conditions [2] . This can be explained by the fact that accumulation of organic acid is strongly influenced by the medium composition [2],[18]. These results show that, in the Aspergillus genus, the major metabolite secreted is different depending on the strain.

Table 1

Highest concentrations of LMWOA and ethanol obtained at day 6 of incubation, for each compound and the corresponding producing strains

Compound	Fungal strain	g.L −1
	Ascomycota
Ethanol	A. niger (BRFM421)	4.1
Gluconic acid	A. niger (BRFM431)	3.7
Formic acid	A. flavipes (BRFM456)	3.3
Citric acid	A. niger (BRFM422)	2.2
Succinic acid	A. flavipes (BRFM456)	1.8
Oxalic acid	A. niger (BRFM420)	1.6
Malic acid	A. niger (BRFM103)	0.6
Acetic acid	A. niger (BRFM428)	0.4
Propionic acid	A. niger (BRFM422)	0.2
Butyric acid	A. flavus (BRFM821)	0.2
Isobutyric acid	A. niger (BRFM432)	0.2
Tartaric acid	A. niger (BRFM420)	0.1
Lactic acid	A. niger (BRFM428)	0.1
Ascorbic acid	A. niger (BRFM280)	<0.05*
Fumaric acid	A. niger (BRFM438)	<0.05*
Itaconic acid	A. terreus (BRFM111)	<0.05*
	Basidiomycota
Ethanol	S. hirsutum BRFM889)	0.2
Gluconic acid	P. coccineus (BRFM1396)	0.6
Formic acid	P. coccineus (BRFM1396)	0.2
Oxalic acid	G. weberianum (BRFM1548)	0.1

*limit of quantification.

Six strains of Aspergillus (BRFM103, BRFM420, BRFM421, BRFM422, BRFM431 and BRFM434) have been selected for further studies due to their high organic acids or ethanol production, and to the variety of organic acids produced. A. brasiliensis BRFM103, A. niger BRFM421, and A. niger BRFM434 were selected for their ability to produce ethanol, 3.6, 4.1, and 2.5 g.L−1, respectively. A. niger BRFM420 was selected for its production of oxalic acid (1.6 g.L−1). A. niger BRFM422 was selected for its production of citric acid (2.2 g.L−1) and A. niger BRFM431 was selected for its production of citric (2.1 g.L−1) and gluconic acids (3.7 g.L−1). At day 6 of incubation 14.2 to 22.9 g.L−1 of glucose remained in the medium and these strains converted 8 to 15% of the glucose consumed to the main organic acid or ethanol (Table 2).

Table 2

Concentrations and conversion yields for the 6 best organic acid producers

	Concentration (g.L −1 )	Mean Y P/S (%)
Aspergillus brasiliensis (BRFM103)
Oxalic acid	0.4 ± 0.1	1.2
Citric acid	0.5 ± 0.1	1.5
Malic acid	0.7 ± 0.2	2.0
Ethanol	3.9 ± 1.0	10.8
Residual glucose	14.2 ± 1.7
Aspergillus niger (BRFM420)
Oxalic acid	2.0 ± 0.4	7.0
Citric acid	0.5 ± 0.1	1.9
Residual glucose	21.6 ± 3.8
Aspergillus niger (BRFM421)
Oxalic acid	0.4 ± 0.1	1.4
Gluconic acid	2.6 ± 0.5	9.5
Malic acid	0.4 ± 0.1	1.6
Ethanol	4.0 ± 0.7	14.5
Residual glucose	22.9 ± 1.1
Aspergillus niger (BRFM422)
Oxalic acid	0.4 ± 0.1	1.3
Citric acid	2.5 ± 0.6	7.7
Tartaric acid	0.2 ± 0.1	0.6
Gluconic acid	3.4 ± 0.1	10.4
Succinic acid	0.8 ± 0.1	2.3
Fumaric acid	0.7 ± 0.2	2.3
Residual glucose	17.6 ± 1.3
Aspergillus niger (BRFM431)
Oxalic acid	0.9 ± 0.1	3
Citric acid	2.4 ± 0.4	8.2
Gluconic acid	4.7 ± 0.6	15.6
Malic acid	0.4 ± 0.1	1.5
Succinic acid	0.7 ± 0.1	2.2
Residual glucose	20.4 ± 1.6
Aspergillus niger (BRFM434)
Oxalic acid	0.6 ± 0.1	2.2
Citric acid	0.4 ± 0.1	1.4
Gluconic acid	2.4 ± 0.8	8.5
Ethanol	2.1 ± 0.5	7.4
Residual glucose	21.9 ± 1.7

Yields are expressed in g of product per g of glucose consumed.

(± SD), n = 3.

In order to confirm the identity of organic acids of applied interest observed by HPLC-UV (i.e. citric, lactic, malic, and oxalic acid), supernatants from fresh cultures of A. brasiliensis BRFM103, A. niger BRFM422 and A. niger BRFM428 were analyzed by GC-MS. These 4 organic acids were detected in all the supernatants. With HPLC-UV, malic acid was not detected in the supernatants of strains BRFM422 and BRFM428, and lactic acid was detected only in BRFM428. This result suggests that these three strains are able to produce citric, lactic, malic and oxalic acid. However, only BRFM103 produced malic acid and BRFM422 produced lactic acid at amounts detectable by HPLC-UV. Besides, we confirm the high ethanol production by BRFM103 and found smaller amounts of ethanol in BRFM422 and BRFM428 as well. This ethanol production was also found by HPLC-RI (data not shown) showing a biological variability compared to the first cultures.

As expected the two main organic acids produced by the 6 Aspergilli strains were citric acid and gluconic acid [2],[18]. Interestingly, all these strains also produced oxalic acid. For most of them the production of oxalic acid was low and ranged from 0.4 to 0.9 g.L−1. This is consistent with the literature since oxalic acid production has been shown to be inhibited at pH below 3 by A. niger [27] and by ammonium and excess of substrate [1],[28]. One exception is A. niger BRFM420 which produced 2 g.L−1 of oxalic and with a conversion yield of 7 g oxalic acid/100 g of glucose consumed. For this strain, the only other organic acid detected was citric acid at 0.5 g.L−1. Even if this conversion yield is low compared to yields obtained in optimum conditions [29], this strain is particularly interesting since it seems more disposed than other Aspergilli to produce oxalic acid, even when grown in conditions not promoting the production of this organic acid.

Regarding ethanol production, the best yield, 14.5%, was observed with A.niger (BRFM421). A. oryzae and Rhizopus oryzae have been shown to convert 51.8% of glucose into ethanol [30]. However, a complex medium was used in this study, therefore glucose was not the sole carbon source. As a comparison, Saccharomyces cerevisiae, the fermentative organism used for industrial ethanol production, has a maximum theorical yield on glucose of 51.1%, and industry processes are considered economically relevant above 90% of this yield [31]. The main drawback of ethanol production from biomass using Saccharomyces is that the naturally occurring yeast cannot metabolize xylose, a product of biomass degradation [32]. These findings could be of interest for the production of 2nd generation ethanol from hemicelluloses and consolidated bioprocessing of biomass to ethanol [33].

Methods

Strains

All the strains were provided by the fungal culture collection (BRFM) of the International Centre of Microbial Resources (CIRM-CF; http://www6.inra.fr/cirm_eng/cirm-cf, Marseille, France) of the French National Institute for Agricultural Research (INRA, Marseille, France). At least one species from each family represented at the CIRM-CF was selected in order to analyze the available biodiversity. More strains and species were studied for families largely represented in the collection to achieve a better geographic diversity (Table 3, Figure 3). In total, 66 strains from 47 different species representing 23 fungal families were studied for organic acid production and other metabolic end-products. 40 of these strains originated from the Ascomycota phylum and 26 strains originated from the Basidiomycota phylum. Strains were cultivated on malt agar medium for mycelium expansion prior to inoculation of liquid cultures.

Table 3

List of the strains studied and their geographic origin and corresponding BRFM numbers

	Current name	Family	Continent	BRFM
Ascomycota
	Cordyceps militaris	Cordycipitaceae	Europe	1581
	Eutypella scoparia	Diatrypaceae	Central America	1014
	Hypocrea lixii	Hypocreaceae	Europe	1058
	Hypocrea lixii	Hypocreaceae	South America	1204
	Hypomyces luteovirens	Hypocreaceae	Europe	1580
	Cosmospora vilior	Nectriaceae	Central America	982
	Nectria pseudotrichia	Nectriaceae	Central America	1017
	Cosmospora vilior	Nectriaceae	Europe	1198
	Haematonectria haematococca	Nectriaceae	South America	1214
	Nectria pseudocinnabarina	Nectriaceae	South America	1287
	Nectria pseudocinnabarina	Nectriaceae	Central America	1288
	Lanatonectria flocculenta	Nectriaceae	South America	1387
	Haematonectria guyanensis	Nectriaceae	South America	1437
	Lasionectria lichenocola	Nectriaceae	Europe	1442
	Sinosphaeria bambusicola	Thyridiaceae	NA*	1245
	Aspergillus brasiliensis	Trichocomaceae	Europe	103
	Aspergillus flavipes	Trichocomaceae	Europe	456
	Aspergillus flavus	Trichocomaceae	NA	99
	Aspergillus flavus	Trichocomaceae	Africa	821
	Aspergillus niger	Trichocomaceae	Africa	107
	Aspergillus niger	Trichocomaceae	NA	280
	Aspergillus niger	Trichocomaceae	Central America	419
	Aspergillus niger	Trichocomaceae	Central America	420
	Aspergillus niger	Trichocomaceae	Central America	421
	Aspergillus niger	Trichocomaceae	Central America	422
	Aspergillus niger	Trichocomaceae	Central America	427
	Aspergillus niger	Trichocomaceae	Central America	428
	Aspergillus niger	Trichocomaceae	Central America	431
	Aspergillus niger	Trichocomaceae	Central America	432
	Aspergillus niger	Trichocomaceae	Central America	434
	Aspergillus niger	Trichocomaceae	Central America	438
	Aspergillus niger	Trichocomaceae	Central America	439
	Aspergillus niger	Trichocomaceae	Europe	449
	Aspergillus oryzae	Trichocomaceae	Europe	488
	Aspergillus tamarii	Trichocomaceae	South America	1520
	Aspergillus terreus	Trichocomaceae	Europe	111
	Aspergillus tubingensis	Trichocomaceae	South America	1521
	Xylaria laevis	Xylariaceae	South America	1243
	Xylaria schweinitzii	Xylariaceae	South America	1447
	Hypoxylon investiens	Xylariaceae	South America	1445
Basidiomycota
	Heterobasidion annosum	Bondarzewiaceae	NA	238
	Gymnopilus junonius	Cortinariaceae	Europe	969
	Ischnoderma benzoinum	Fomitopsidaceae	Europe	1133
	Postia stiptica	Fomitopsidaceae	Europe	1148
	Amauroderma sp.	Ganodermataceae	South America	1359
	Ganoderma weberianum	Ganodermataceae	South America	1548
	Gloeophyllum sepiarium	Gloeophyllaceae	Europe	988
	Fomitiporia mediterranea	Hymenochaetaceae	Europe	1315
	Dichostereum effuscatum	Lachnocladiaceae	Europe	91
	Lentinula edodes	Marasmiaceae	NA	353
	Grifola frondosa	Meripilaceae	Europe	1162
	Abortiporus biennis	Meruliaceae	Europe	1215
	Omphalotus olearius	Omphalotaceae	Europe	1195
	Phanerochaete chrysosporium	Phanerochaetaceae	Europe	413
	Pleurotus ostreatus	Pleurotaceae	Europe	1326
	Grammothele fuligo	Polyporaceae	South America	1046
	Daedaleopsis confragosa	Polyporaceae	Europe	1187
	Perenniporia ochroleuca	Polyporaceae	Europe	1192
	Earliella scabrosa	Polyporaceae	South America	1220
	Tinctoporellus epimiltinus	Polyporaceae	South America	1229
	Trametes menziesii	Polyporaceae	Oceania	1281
	Trametes sp.	Polyporaceae	South America	1361
	Pycnoporus coccineus	Polyporaceae	Oceania	1396
	Polyporus squamosus	Polyporaceae	Europe	1531
	Stereum hirsutum	Stereaceae	Europe	889
	Panellus serotinus	Tricholomataceae	Europe	1284

Families are sorted in alphabetical order, when several strains from one family were tested; species were sorted in alphabetical order and by increasing BRFM number for strains from the same species.

*NA: not available.

Figure 3

Repartition of the strains selected for the screening in the and phyla. 40 strains of Ascomycota representing 6 families and 26 strains of Basidiomycota representing 16 families were screened.

Chemicals and reagents

Ultrapure water (conductivity 18.2 mΩ) was used in all experiments. For fungal cultures, malt extract was purchased from VWR, EDTA-Na2 and glucose were from Sigma, (NH4)2SO4 and CaCl2 were from Panreac, H3BO3, MnCl2, FeSO4, CuSO4, CoCl2 Na2MoO4, MgSO4, KH2PO4, and ZnSO4 were purchased from Prolabo and Bacto agar was purchased from Fischer.

For analytic methods, CD3OH, HPLC grade organic acid standards (acetic, adipic acid, L-ascorbic acid, benzoic acid, butyric acid, citric acid, isobutyric acid, formic acid, fumaric acid, L-(+)-lactic acid, DL-isocitric acid trisodium salt hydrate, maleic acid, malonic acid, D-(+)-malic acid, oxalic acid, phytic acid, propionic acid, (−)quinic, succinic acid, shikimic acid,D-(−)-tartaric acid), methylchloroformate (MCF), and glucose were purchased from Sigma, HPLC grade ethanol was purchased from Fluka. Dichloromethane was from Carlo Erba Reagents.

Liquid cultures

The liquid medium was composed of (NH4)2SO4 (8 mM), CaCl2 (1 mM), KH2PO4 (11 mM), MgSO4 (2 mM), glucose 5% (wt/vol) and trace elements and had an initial pH of 5.5. The final concentrations of trace elements was ZnSO4 (76 μM), H3BO3 (178 μM), MnCl2 (25 μM), FeSO4 (18 μM), CoCl2 (7.1 μM), CuSO4 (6.4.μM), Na2MoO4 (6.2 μM), and EDTA-Na2 (174 μM). The glucose concentration had to be high, in order to be close to industrial conditions for organic acid production in Ascomycota [34]. In previous screenings of Basidiomycota for organic acid production, the glucose concentration was set to 5% [12]. This concentration was therefore chosen to be suitable for both Ascomycota and Basidiomycota growth. For Basidiomycota, which hardly grow on such media, Tatum vitamins [35] and yeast extract (0.03 g.L−1) were added to the liquid medium. Cultivations were carried out in 250 mL baffled flasks to facilitate oxygen transfert. They contained 100 mL of liquid medium and were incubated for 6 days at 30°C in an orbital incubator at 120 and 140 rpm for Ascomycota and Basidiomycota, respectively. For Aspergilli, the liquid medium was inoculated at an initial titer of 2 × 106 spores.mL−1. For other fungi (Basidiomycota and other Ascomycota), that do not produce enough spores in our growth conditions, fungal disks, 4 mm in diameter, were collected from the solid medium. For each strain, 3 tubes containing 1 fungal disk and 1 mL sterile water were crushed at a frequency of 4 s−1 during 60 s using a FastPrep-24 (MPBio, Solon, OH, USA). The 3 tubes were then mixed together. Afterwards, 1 mL of the inoculum preparation was added to each flask. Cultures were carried out in triplicates.

Analytical methods

During the incubation, the acidity of the culture media was evaluated daily with pH paper (Duotest®, Machery-Nagel, Düren, Germany). At days 3 and 6 of incubation, 2 mL samples were harvested from the media. Mycelium was removed by centrifugation and the supernatant was collected after ultra-filtered using Vivaspin® 5kD (VWR, Strasbourg, France) tubes to remove proteins. The filtrates were analyzed for organic acids by HPLC (Agilent 1100 series HPLC, Santa Clara, CA, USA) using an Aminex HPX-87H organic acid analysis column (100 mm × 7.8 mm, Biorad, Marnes-la-Coquette, France). The column was equilibrated in 2.5 mM H2SO4 at 35°C and samples were eluted with 2.5 mM H2SO4 at a 0.6 mL.min−1 flow rate. Organic acids were detected with a UV detector at 210 nm (G1314A, Agilent HPLC 1100 series) and ethanol and glucose were detected with a differential refractometer (HP1047A, Hewlett Packard). Data were acquired with ChemStation software (Agilent, Hewlett Packard, Waldbronn, Germany). The first analysis of metabolite secretion was performed with pools of the three replicates of each strain in order to get an estimation of the mean organic acids production. The supernatants of the higher producers were then analyzed separately and the identification of organic acids and ethanol was further studied using GC-MS.

Supernatants of the cultures were harvested by centrifugation and organic acids were directly derivatized, without ultrafiltration, using MCF as previously described [36], with some modifications. Briefly, 190 μL of the supernatants were directly alkalinized with 10 μL NaOH 2.5 M and derivatized by two consecutive additions of 20 μL MCF. After derivatization, the methylated compounds were extracted with dichloromethane. GC-MS analysis of organic acids was performed with an Agilent 5973 N system, equipped with an Omegawax (Supelco, Bellefont, PA, USA), 30 m × 250 μm i.d. × 0,25 μm thick films. The carrier gas was helium at 35 cm.s−1. The oven program temperature started at 40°C during 3 min, then rose at 8°C per minute to 230°C and held at this temperature for 15 min. 2 μL of extract were injected in split injector port with split ratio of 10. Mass spectra in the 29 to 400 m/z range were recorded at a scanning speed of 2 scans.s−1 and an electronic ionization at 70 eV. Compounds were identified by matching compound mass spectra to the NIST library and using pure authentic chemical standards for each organic acid studied, derivatized using the same process as biological samples.

Ethanol concentration was determined in 2 mL supernatants by Head-space-GC-MS with addition of 0.4 mL of the internal standard CD3OH at 0.6 g.L−1. A calibration curve was prepared with 201.8, 403.6 and 807.2 μg of ethanol with CD3OH as internal standard as above. For GC we used the GC–MS QP2010 Shimadzu with capillary column Cp_wax_52cb 30 m × 0.32 mm × 0.5 μm (Varian, Inc, Palo Alto, USA) equipped with an autosampler AOC5000. The sealed vials were placed at 50°C for 8 min with 500 rpm shaking before 0.5 mL of the headspace were drawn out with a gas syringe heated at 60°C and injected with in a split injector with a split ratio of 10. The carrier gas was helium at 35 cm.s−1 and oven temperature was isothermal at 50°C. Mass detector conditions were: electronic impact ionization mode (70 eV), temperature of source 200°C with data collected using SIM for selected ions m/z 45/46 and 35/30 for ethanol and CD3OH respectively.

Conclusion

The potentiality of a wide panel of fungus for organic acids production has been studied in glucose based liquid media at acidic pH to take into account industrial up- and down-stream technical and economical issues. Strains were sorted in two clusters considering their organic acid and ethanol production at day 6 of incubation, showing that some strains, even from the same species, seem to have particular predispositions for some metabolites.

Among 26 Basidiomycota tested, only two: Postia stiptica (brown-rot) and Ganoderma weberianum (white-rot) produced oxalic acid. Ethanol is the common metabolite in the fungal kingdom, regardless the geographic origin of the strains, but with different extent depending on the strain. Although yeasts have very competitive ethanol productivity on simple sugars, our best ethanol producers may be good candidates for consolidated bioprocessing (CBP) of cellulosic biomass for second generation ethanol production.

Among the Ascomycota, Aspergilli clearly make a distinct cluster for their various and high concentration organic acid production; this illustrates the relevance of organic acids in the chemotaxonomy of Aspergilli. Some of these strains showed particular ability to produce malic, oxalic, gluconic and citric acids or ethanol at low pH. This production could be further improved by genetic modifications. The high intra-specific variability in metabolite production stresses the importance of screenings for a good choice of studied strains.

This study provides a better knowledge of the capability of filamentous fungi to produce organic acids which should allow a greater exploitation of filamentous fungi in synthetic biology, metabolic studies and industrial exploitation of organic acids.