Utilization and optimization of a waste stream cellulose culture medium for pigment production by Penicillium spp.



James Smith, QUT, Science and Engineering Faculty, School of Earth, Environmental and Biological Sciences, GPO Box 2434, Brisbane QLD 4001, Australia. E-mail: smithjj16@tpg.com.au



This research sought to determine optimal corn waste stream–based fermentation medium C and N sources and incubation time to maximize pigment production by an indigenous Indonesian Penicillium spp., as well as to assess pigment pH stability.

Methods and Results

A Penicillium spp. was isolated from Indonesian soil, identified as Penicillium resticulosum, and used to test the effects of carbon and nitrogen type and concentrations, medium pH, incubation period and furfural on biomass and pigment yield (PY) in a waste corncob hydrolysate basal medium. Maximum red PY (497·03 ± 55·13 mg l−1) was obtained with a 21 : 1 C : N ratio, pH 5·5–6·0; yeast extract-, NH4NO3-, NaNO3-, MgSO4·7H2O-, xylose- or carboxymethylcellulose (CMC)-supplemented medium and 12 days (25°C, 60–70% relative humidity, dark) incubation. C source, C, N and furfural concentration, medium pH and incubation period all influenced biomass and PY. Pigment was pH 2–9 stable.


Penicillium resticulosum demonstrated microbial pH-stable-pigment production potential using a xylose or CMC and N source, supplemented waste stream cellulose culture medium.

Significance and Impact of the Study

Corn derived, waste stream cellulose can be used as a culture medium for fungal pigment production. Such application provides a process for agricultural waste stream resource reuse for production of compounds in increasing demand.


Naturally derived colourants appear set to overtake synthetic colourants in total market value as manufacturer demand rises for naturally derived ingredients, particularly in food applications (Mapari et al. 2009). Naturally derived pigment use has a predicted annual growth rate of 5–10%, while synthetic dye colourants are forecasted to grow at 3–5% (Downham and Collins 2000). The value of the 2007 international food colourant market alone was estimated at $1·15 billion USD, increasing 2·5% from 2004 (Mapari et al. 2010).

Environmental and human toxicity issues associated with synthetic pigments and their production have also increased interest in natural pigment use (Venil and Lakshmanaperumalsamy 2009). Existing authorized natural food colourants are of either plant or animal origin, production of which is subject to variable agricultural, seasonal and climatic conditions. In Indonesia, plant-derived red pigments from the roselle (Hibiscus sabdariffa L) and achiote (Bixa orellana) have been used as food colourants, but are only seasonally available.

Recent research has focused on developing environmentally friendly pigment production methods, particularly those utilizing microbial biosynthesis. Microbially derived colourants play a significant role as food colouring agents, due to simplified downstream processing and lack of impact of agricultural, climatic and seasonal conditions on production (Vidyalakshmi et al. 2009). A number of micro-organisms have the ability to produce pigments at high yields. Among these, fungal species have attracted substantial consideration due to their ability to produce pigments of differing toxicity and chemical structure (Hajjaj et al. 1999).

Pigments are used in the food industry as colourants, additives, colour intensifiers and antioxidants (Mendez et al. 2011). Isolation and identification of new pigment producing fungal strains, use of inexpensive and/or waste feedstock and fermentation process development are of particular interest to maximize pigment yields (PYs). The diversity of fungal pigments in terms of chemical structure and colour range may add new or additional hues to the colour palette of existing, naturally derived pigment colourants (Mapari et al. 2006). Many Penicillium spp. produce pigments, but only Penicillium oxalicum has been used on an industrial scale to produce food colourant (Mapari et al. 2006; Dufossé 2006; Mapari et al. 2009). Coculture of Penicillium sp. HSD07B and Candida tropicalis on a glucose-potato-based medium produced a pH-stable, red pigment showing no acute toxicity at a yield of 2·75 and 7·7 g l−1 in shake flask culture and a 15-l bioreactor, respectively (Hailei et al. 2011). However, other than this latter study, Penicillium spp. pigment production has rarely used cellulose as a carbon source, despite cellulose availability as an agricultural waste product.

Lignocellulose is the most abundant renewable, photosynthetically produced biomass, with a global annual production of approximately 200 billion metric tons (Ragauskas et al. 2006; Zhang et al. 2006). Corn (Zea mays L.) is a primary global food crop, with relatively high production (as kernels) in Indonesia compared to other food crops, at approximately 13 million tons per annum (2007). Corncobs are a lignocellulosic waste product representing up to 30% of corn by weight. There is little productive use of waste corncobs in Indonesia, with most discarded or burned. Corncobs can be used as an alternative livestock feed, primarily during the dry season. However, corncob feed is very high in crude fibre. The agricultural by-product lignocellulosic materials consist of lignin, hemicelluloses and celluloses. The lignin fraction is a noncarbohydrate amorphous organic compound, consisting of condensed phenolic units (Chang et al. 1981).

Hydrolysis of lignocelluloses to release glucose is the most important step in the economic production and utilization of renewable cellulosic materials. Although such raw materials are inexpensive, pretreatment is generally required to improve subsequent utilization of lignocellulosic materials. Bioconversions require hydrolysis of lignocellulose polymers to their constituent monomeric sugars prior to microbial fermentation, with lignocellulose hydrolysis generally achieved using either acid (Lee et al. 2000) and/or enzymatic treatment(s) (Sreenath and Jeffries 2000). Acid hydrolysis is a simple, inexpensive method. However, fermentative conversion of lignocellulose hydrolysate can be severely inhibited by some lignocellulose hydrolysis by-products such as furfural (Olsson and Hähen-Hägerdal 1996; Larsson et al. 2001), which has been shown to have inhibitory effects on yeast-specific growth and fermentation rates (Palmqvist et al. 1999; Tofighi et al. 2010). In this study, we isolated an indigenous Indonesian red pigment producing Penicillium spp., and through C, N and furfural supplementation determined the optimal corncob waste stream material-based culture medium and incubation time for pigment and fungal biomass production. We also examined the impact of the presumed end product inhibitor furfural on biomass and pigment production.

Materials and methods

Fungal isolation and identification

Soil samples were collected from three locations at Baluran National Park (7°51′53″S; 114°21′32″E) East Java, Indonesia, and transported to the laboratory in sterile plastic bags. Samples were serially diluted (10−6) in sterile distilled water, spread plated onto potato dextrose agar (PDA) and incubated at 25°C for 3 days. All plate and flask incubations in this study were carried out at 60–70% relative humidity (r.h.). Presumptive Penicillium spp. were identified by mycelium colour and microscopic morphological examination. These were subcultured to Czapek yeast autolysate (CYA) and malt extract agars (MEA) (25°C, 3 day). Presumptive pigment producing Penicillium spp. were selected by mycelium pigmentation, characteristic zonation, pigment surrounding mycelia and reverse-view colour under both illuminated (fluorescent light) and dark incubation conditions. Confirmation as Penicillium resticulosum was performed via microscopic examination of morphological characteristics using light microscopy at 400–800× magnification of wet mounts stained with 1% lactophenol cotton blue and standard reference identification keys (Pitt 1973; Watanabe 1975; Samson and Pitt 1985). The isolate selected for use in this study was chosen based on visually apparent maximum red pigment colour production on PDA. The master stock culture was kept on PDA at 25°C and subcultured every 3 months.

Corncob acid hydrolysis

Locally farm-sourced corncobs from Surabaya, Indonesia, were air-dried then ground to approximately 2-mm-diameter particles using a mill. The modified method of Oliveira et al. (2006) was used to prepare acid hydrolysates for subsequent fermentation. Briefly, 1-kg dry milled corncob was mixed with 10 l, 2·5% H2SO4, autoclaved (15 min, 121°C autoclave procedure used for all media this study), cooled to room temperature, the liquid fraction aseptically filtered through sterile cotton cloth and filtrate pH adjusted to 6·5 with 1 mol l−1 NaOH. Pb acetate and potassium oxalate were then added to final concentrations of 0·1% each and the filtrate refiltered (Whatman No 1 paper). Filtrate was stored at 1–5°C in the dark until used.

Culture and inoculum preparation

Working stock cultures were prepared from stock, 7 days, 25°C PDA plate cultures subcultured from the master stock. Conidia were aseptically sampled by scraping the top of a mycelium with an inoculating loop and transfer to 10-ml sterile water. Inoculum stock suspension was prepared from working stock and diluted to 1·7 × 106conidia ml−1, as enumerated using a haemocytometer.

Carbon and nitrogen source

The basal medium contained 1000-ml corncob hydrolysate, 5·0 mg l−1 KH2PO4, and 0·02 g l−1 MgSO4·7H2O. This was supplemented with carbon [xylose, glucose, sucrose, carboxymethylcellulose (CMC)] and nitrogen (peptone, yeast extract, NH4NO3, NaNO3) to final individual concentrations of 20 g l−1, with additional unsupplemented controls. Media were mixed thoroughly and 50 ml individually dispensed into five 250-ml Erlenmeyer flasks, autoclaved, cooled to room temperature, individually inoculated with 1 ml of inoculum stock suspension and incubated at 28–29°C, 50% r.h. in the dark for 12 days in a rotary incubator at 60 rev min−1. This inoculation and incubation method was used for all cultivation in this study.

Furfural supplementation

Liquid basal medium (1500 ml) containing 20·0 g l−1 glucose and 3·0 g l−1 yeast extract was dispensed into six 250-ml Erlenmeyer flasks, autoclaved and cooled to room temperature. Furfural was added to individual flasks to final concentrations of 0·0, 1·0, 2·0, 3·0, 4·0 and 5·0 g l−1. Fifty millilitre was then aseptically dispensed into individual five Erlenmeyer flasks, 1-ml inoculum stock suspension added and flasks incubated as described above.

Cellulose and nitrogen concentration

Five cellulose and nitrogen concentrations (0, 10·0, 20·0, 30·0 and 40·0 g l−1 each, respectively) in basal medium were used to examine their effects on biomass and PYs. Nitrogen source was a mixture of yeast extract (3 g l−1), NH4NO3 (4 g l−1) and NaNO3 (3 g l−1). Cellulose and nitrogen sources were added to basal medium in 250-ml Erlenmeyer flasks, mixed thoroughly, autoclaved and cooled to room temperature. Flasks were inoculated with 1-ml inoculum stock suspension and incubated as described above.

Initial medium pH

To examine the effect of initial medium pH, 50-ml basal medium containing 20 g l−1 CMC, 3·0 g l−1 yeast extract, 4·0 g l−1 NH4NO3 and 3·0 g l−1 NaNO3 was aliquoted into 5, 250-ml Erlenmeyer flasks and the pH of each adjusted to 3·5, 4·5, 5·5, 6·5 and 7·5 prior to autoclaving. After cooling to room temperature, flasks were inoculated with 1-ml inoculum stock suspension and incubated as described above.

Incubation period

Effect of incubation period on biomass and PYs was examined using a basal medium supplemented with 30·0 g l−1 CMC (CMC amount found to produce maximum pigment), 3·0 g l−1 yeast extract, 4·0 g l−1 NH4NO3 and 3·0 g l−1 NaNO3. Erlenmeyer flasks (250 ml) containing 50-ml sterile medium were inoculated with 1-ml inoculum stock suspension and incubated as described above. Dry biomass and PY analyses were carried out every 3 days up to 21-days incubation. All data presented are means of three simultaneously incubated fermentation culture replicates.

Dry biomass

To determine dry biomass, whole flask cultures were poured through predried (100°C) preweighed Whatman No.1 filter paper. Retained mycelial material was washed with distilled water then ethanol until colourless and dried at 100°C to constant weight (48 h).

Pigment yield and pH stability

Culture filtrates were centrifuged at 3000 g, 15 min. Supernatant pigment (approximately 50 ml) was transferred to a 200-ml-capacity borosilicate glass tube and an equal volume of ethyl acetate added. The pH was adjusted to 3·0 using 2 mol l−1 HCl, the sample mixed vigorously and left to stand for 15 min. The upper ethyl acetate layer was removed and evaporated to a viscous suspension using a rotary vacuum evaporator (Heidolph VV2011; Heidolph Instruments Labortechnik, Schwabach, Germany) at 50°C. The resulting pigment extract viscous suspension was transferred to a preweighed watch glass and dried to constant weight at 50°C. Dried pigment (10 mg) was dissolved in 10-ml spectrophotometric grade ethyl acetate and 0·1 ml of this pigment solution added to individual 50-ml aliquots of ethyl acetate in 100-ml Erlenmeyer flasks pre-adjusted to eight different respective pH levels (2, 3, 4, 5, 6, 7, 8 and 9) with 0·1 mol l−1 HCl or NaOH, and solutions allowed to stand for 1 h at room temperature under ambient fluorescent light illumination. Each pigment solution was then filtered through a 1·5-μm-pore size glass fibre filter and filtrate visible light and UV absorption spectra (200–700 nm) measured using a Shimadzu 1700 scanning spectrometer (used for all spectrophotometric measurements in this study).

Determination of glucose consumption

Glucose concentration was determined using the dinitrosalicylic acid (DNSA) method (AOAC 1990; method 996.11). Two millilitre of initial, nonwater-rinsed liquid sample filtrate from fermentations (see dry biomass determination above) was placed in a glass test tube, 1-g activated charcoal added, the mixture shaken thoroughly and filtered through Whatman No. 1 filter paper. One-millilitre of filtrate was placed in a test tube, two drops alkaline DNSA added, the tube placed in boiling water for 5 min, allowed to cool to room temperature and absorbance measured at 540 nm. Glucose was measured after at 0, 24, 48, 72, 96, 120, 144 and 168 h of fermentation.

Determination of organic carbon

Levels of total organic carbon (TOC) were determined using the wet oxidation method of Walkey and Black (1965). One hundred millilitre of liquid culture was evaporated at 100°C for approximately 2 h to obtain a dried powder, 0·5 g of which was used for TOC determination.

Nitrogen determination

Nitrogen (NH4-N) concentration was determined using the method of the American Society of Agronomy and Soil Science Society of America (1982). Ten-millilitre culture medium was evaporated at 100°C for approximately 2 h to obtain a dried powder. Samples (50 mg) were added to digestion tubes and 1-g selenium mixture (mashed 1·55 g CuSO4, 96·9 g Na2SO4 and 1·55 g selenium) and 3-ml 97% H2SO4 added, mixed and digested at 350°C for 4 h to obtain a colourless extract, cooled to room temperature, diluted to 50 ml with distilled water, shaken vigorously and left to stand overnight. Two-millilitre of extract was placed, transferred to a new borosilicate glass test tube and 4-ml tartrate buffer (50 g NaOH and 50 g KNaC4H4O6 l−1) and sodium phenate solution (100 g NaOH and 125 g phenol l−1) successively added, mixed and let stand for 10 min. Four-millilitre, 5% NaOCl was then added, the mixture shaken, let stand for 10 min and absorbance measured at 636 nm. (NH4)2SO4 was used to prepare N standards.

Statistical analysis

Tukey's honestly significant difference multiple comparison tests were used to segregate significantly different treatments using SPSS 13 software. Analysis of variance (anova) was performed to determine significant differences between experiments (P < 0·05).


Penicillium isolate morphology

A total of six soil samples were taken from three locations in Baluran National Park, Indonesia. Nine Penicillium sp. isolates were obtained, one of which produced visible red pigmentation on PDA, MEA and CYA when incubated in the dark only (Fig. 1). Pigment production visually appeared greater on PDA and MEA than CYA, and growth on PDA under lighted conditions showed very faint red pigmentation (Fig. 1a–f). Colour surrounding mycelia on PDA medium suggested pigment was excreted and diffusible (Fig. 1a). Macroscopic and microscopic morphology of the Penicillium sp. isolate are presented in Table 1 and Fig. 1. The isolate was identified as P. resticulosum and designated strain Blr1 based on classical taxonomic criteria.

Table 1. Macroscopic and microscopic characteristic of Penicillium resticulosum Blr1 grown at 25°C for 7 days at 60–70% relative humidity
1MacroscopicPotato dextrose agar: good growth, velvety, zonate white-green-white-green; colony diameter 27·0–30·0 mm (Fig 2a), reverse side; red zone 31–32 mm in diameter (Fig 2b).
Malt extract agar: grew rapidly, top side zonate white-red-white, colony diameter 27·0–32·0 mm (Fig 2c), reverse side; red zone 25·5–26·0 mm in diameter (Fig 2d), pigment not produced under lighted incubation conditions (Fig 2g,h).
Czapek yeast autolysate: grew more slowly than malt extract agar, top side zonate white-green-white, colony diameter 24·0–26·0 mm (Fig 2e), reverse side; red zone 10·1–11·5 mm in diameter (Fig 2f), pigment not produced under lighted incubation conditions (Fig 2i,j).
2MicroscopicConidiophores 60–130 μm tall, with 10–13·8 × 2·5–2·8 μm primary branches were hyaline, erect, branched with 2–3 metula, 3–4 verticillate phialides and catevulate conidia in each phialide, forming rather compact cylindrical grey-green conidial head, conidial heads and conidial head lancet. Conidia 2·1–2·8 μm in diameter were pale green to dark-brown in mass, subglobose and minutely echinulate. Conidiophores and hyphae were septate generative and vegetative.
Figure 1.

Penicillium resticulosum Blr1 mycelia grown at 25°C for 3 days under fluorescently lit conditions in respective obverse/reverse views on potato dextrose agar (a)/(b), malt extract agar (MEA) (c)/(d), Czapek yeast autolysate (CYA) (e)/(f) and under dark incubation conditions in respective obverse/reverse views on MEA (g)/(h) and CYA (i)/(j). Arrows indicate diffuse (a) and mycelium-associated (b, c, d, f) pigment.

Pigment pH stability

There was a slight reduction in absorbance at the absorbance maximum (λmax) (496 nm) in ethyl acetate at lower pH values (pH 2, 3), but overall pigment spectra were consistent with varying pH (Fig. 2). Absorbance maxima at pH 2, 3, 4, 5, 6, 7, 8 and 9 were 0·33, 0·35, 0·36, 0·36, 0·37, 0·37, 0·37, 0·37 and 0·38, respectively. This represented λmax reductions from pH 7 of 10·81, 5·41 and 2·70% at pH 2·0, 3·0 and 4·0, respectively.

Figure 2.

Ultraviolet/visible light spectra of pigment from Penicillium resticulosum Blr1 in ethyl acetate stored at pH 2–9 for 1 h under fluorescent light illumination at room temperature. image, ethyl acetate; image, pH 2·0; image, pH 3·0; image, pH 4·0; image, pH 5·0; image, pH 6·0; image, pH 7·0; image, pH 8·0; image, pH 9·0.

Effect of carbon sources

Carbon source type had a significant (P < 0·05) effect on biomass dry weight (BDW) and PY as shown in Fig. 3. BDW with different media varied between 0·86 ± 0·12 and 3·32 ± 0·44 g l−1, while PY varied between 0·12 ± 0·03 and 0·55 ± 0·07 g l−1. Carbon source addition (glucose, sucrose, xylose and CMC) to the corncob hydrolysate basal medium produced a significant (< 0·05) increase in both BDW and PY. BDW in the basal medium plus xylose (3·32 ± 0·55 g l−1) or CMC (3·26 ± 0·54 g l−1) was significantly (< 0·05) higher than that in the basal medium alone (0·86 ± 0·12 g l−1), basal medium plus glucose (2·08 ± 0·31 g l−1) or sucrose (1·93 ± 0·51 g l−1), but not significantly (P > 0·05) different in BDW for xylose or CMC. However, PY in the basal medium plus xylose (0·55 ± 0·07 g l−1) or CMC (0·52 ± 0·05 g l−1) was significantly (< 0·05) higher than the basal medium alone (0·12 ± 0·03 g l−1), basal medium plus glucose (0·21 ± 0·07 g l−1) or sucrose (0·22 ± 0·06 g l−1), but no significant (P > 0·05) difference was observed in PY between basal medium alone or plus xylose or CMC.

Figure 3.

Effect of different carbon source supplementation on biomass dry weight and pigment yield (PY) by Penicillium resticulosum Blr1 in corncob hydrolysate basal medium. BM, basal medium; CMC, carboxymethylcellulose. Values and error bars represent means ± SD (= 5). anova was followed by Tukey's test. a, b, c < 0·05 and *, **, *** < 0·05 within respective groups. image, PY; image, biomass.

Effect of furfural

Addition of 1–5 g l−1 furfural to the growth medium significantly (< 0·05) decreased glucose consumption, BDW and PY of P. resticulosum Blr1 (Fig. 4a,b). Increasing amounts of furfural (1, 2, 3, 4 and 5 g l−1) in the medium progressively decreased glucose consumption to 66·2 ± 0·3; 55·5 ± 0·1; 48·1 ± 0·4; 42·3 ± 0·2 and 32·3 ± 0·3%, respectively, of that in furfural-free medium (Fig. 4a). This indicated a furfural-concentration-dependent suppression of glucose uptake. While mean BDW in furfural-free medium was 5·49 ± 0·24 g l−1, furfural progressively reduced BDW with increasing furfural concentration to 4·83 ± 0·93; 4·29 ± 0·39; 3·80 ± 0·33; 3·31 ± 0·52; and 2·52 ± 0·75 g l−1, at 1, 2, 3, 4 and 5 g l−1 furfural, respectively. Furfural also progressively reduced PY with increasing furfural concentration from 0·61 ± 0·02 g l−1 in furfural-free medium to 0·51 ± 0·13; 0·45 ± 0·02, 0·33 ± 0·09; 0·23 ± 0·09 and 0·16 ± 0·03 g l−1 at 1, 2, 3, 4 and 5 g l−1 furfural, respectively.

Figure 4.

Effect of furfural addition on glucose consumption (a) and biomass dry weight and pigment yield (PY) (b) by Penicillium resticulosum Blr1 in corncob hydrolysate basal medium supplemented with glucose and yeast extract. Values and error bars represent means ± SD (= 5). anova was followed by Tukey's test with a, b, c, d, e, f < 0·05; and *, **, ***, #, ##, ### < 0·05 within respective groups. image, PY; image, biomass.

Effect of nitrogen sources

Addition of nitrogen to the basal medium significantly (< 0·05) increased both BDW and PY. BDW in the basal medium alone (0·86 ± 0·22 g l−1) was significantly (< 0·05) lower than that in the basal medium plus NaNO3 (2·76 ± 0·56 g l−1), NH4NO3 (3·36±0·19 g l−1), yeast extract (3·78 ± 0·43 g l−1) and peptone (4·04 ± 0·54 g l−1). PY in the basal medium (0·25 ± 0·03 g l−1 substrate) was also significantly (< 0·05) lower than that in the basal medium plus NaNO3 (0·34 ± 0·04 g l−1), NH4NO3 (0·35 ± 0·03 g l−1), yeast extract (0·35 ± 0·02 g l−1) and peptone (0·36 ± 0·04 g l−1). However, no significant (> 0·05) difference in BDW or PY was observed between nitrogen source type (Fig. 5).

Figure 5.

Effect of different nitrogen source types in corncob hydrolysate culture medium on Penicillium resticulosum Blr1 biomass dry weight and pigment yield (PY). BM: basal medium; NO3: NaNO3; NH4: NH4NO3; YE: yeast extract; image: PY; image: biomass. Values and error bars represent means ± SD (= 5). anova was followed by Tukey's test. a, b, c < 0·05; and *, ** < 0·05 within respectively groups.

Effect of carbon and nitrogen concentration

Carbon and nitrogen concentration significantly (< 0·05) affected BDW and PY of (Table. 2). Maximum BDW and PY were obtained in corncob hydrolysate medium supplemented with 30·0 g l−1 CMC, 3·0 g l−1 NaNO3, 4·0 g l−1 NH4NO3, and 3·0 g l−1 yeast extract, 5·0 mg l−1 KH2PO4 and 0·02 g l−1 MgSO4·7H2O with a C : N ratio of 21. C : N ratios higher or lower than 21 decreased both BDW and PY.

Table 2. Effect of different cellulose and nitrogen concentrations on fermentation biomass dry weight and pigment yield of Penicillium resticulosum Blr1 in waste corncob hydrolysate basal medium
NoMedium compositionC : N ratioBiomass dry weight (g l−1)Pigment yield (g l−1)
  1. BM, basal medium; YE, yeast extract; CMC, carboxymethylcellulose.

  2. Values are means ± SD (= 5) anova followed by Tukey's test. a, b and c designate groups for between which anova < 0·05.

1BM (basal medium)20 : 11·13 ± 0·60a0·22 ± 0·05a
Effect addition of carbon
2BM + 10 g l−1 CMC25 : 11·07 ± 0·10a0·15 ± 0·12a
3BM + 20 g l−1 CMC27 : 11·22 ± 0·10a0·17 ± 0·05a
4BM + 30 g l−1 CMC28 : 12·19 ± 1·00b0·18 ± 0·06a
5BM + 40 g l−1 CMC30 : 12·12 ± 0·20a0·21 ± 0·02a
Effect addition of nitrogen
6BM + 3·0 g l−1 YE + 4·0 g l−1 NH4NO3 + 3·0 g l−1 NaNO316 : 11·55 ± 0·10a0·19 ± 0·06a
7BM + 6·0 g l−1 YE + 8·0 g l−1 NH4NO3 + 6·0 g l−1 NaNO313 : 11·44 ± 0·70a0·17 ± 0·13a
8BM +  9·0 g l−1 YE + 12·0 g l−1 NH4NO3 + 9·0 g l−1 NaNO311 : 11·19 ± 0·20a0·15 ± 0·10a
9BM + 12·0 g l−1 YE + 16·0 g l−1 NH4NO3 + 12·0 g l−1 NaNO39 : 11·18 ± 0·60a0·15 ± 0·12a
Effect addition of carbon and nitrogen combination
10BM + 10 g l−1 CMC + 3·0 g l−1 YE + 4·0 g l−1 NH4NO3 + 3·0 g l−1 NaNO319 : 11·40 ± 0·40a0·19 ± 0·08ab
11BM + 10·0 g l−1 CMC + 6·0 g l−1 YE + 8·0 g l−1 NH4NO3 + 6·0 g l−1 NaNO315 : 12·32 ± 0·90a0·25 ± 0·04b
12BM + 10 g l−1 CMC + 9·0 g l−1 YE + 12·0 g l−1 NH4NO3 + 9·0 g l−1 NaNO313 : 13·24 ± 1·00b0·17 ± 0·11a
13BM + 10 g l−1 CMC + 12·0 g l−1 YE + 16·0 g l−1 NH4NO3 + 12·0 g l−1 NaNO311 : 12·64 ± 0·60ab0·14 ± 0·07a
14BM + 20 g l−1 CMC + 3·0 g l−1 YE + 4·0 g l−1 NH4NO3 + 3·0 g l−1 NaNO320 : 12·97 ± 1·00c0·30 ± 0·02b
15BM + 20·0 g l−1 CMC + 6·0 g l−1 YE + 8·0 g l−1 NH4NO3 + 6·0 g l−1 NaNO316 : 12·87 ± 0·60bc0·21 ± 0·05b
16BM + 20 g l−1 CMC + 9·0 g l−1 YE  + 12·0 g l−1 NH4NO3 + 9·0 g l−1 NaNO314 : 12·86 ± 0·80bc0·21 ± 0·05b
17BM + 20 g l−1 CMC + 12·0 g l−1 YE + 16·0 g l−1 NH4NO3 + 12·0 g l−1 NaNO311 : 12·19 ± 1·00b0·18 ± 0·06a
18BM + 30 g l−1 CMC + 3·0 g l−1 YE + 4·0 g l−1 NH4NO3 + 3·0 g l−1 NaNO321 : 13·55 ± 0·60c0·50 ± 0·06c
19BM + 30·0 g l−1 CMC + 6·0 g l−1 YE + 8·0 g l−1 NH4NO3 + 6·0 g l−1 NaNO317 : 12·66 ±  0·50b0·30 ± 0·01b
20BM + 30 g l−1 CMC + 9·0 g l−1 YE + 12·0 g l−1 NH4NO3 + 9·0 g l−1 NaNO314 : 12·18 ± 0·50ab0·29 ± 0·02b
21BM + 30 g l−1 CMC + 12·0 g l−1 YE + 16·0 g l−1 NH4NO3 + 12·0 g l−1 NaNO312 : 11·85 ± 0·60a0·17 ± 0·02a
22BM + 40 g l−1 CMC + 3·0 g l−1 YE + 4·0 g l−1 NH4NO3 + 3·0 g l−1 NaNO323 : 13·42 ± 1·00c0·32 ± 0·01b
23BM + 40·0 g l−1 CMC + 6·0 g l−1 YE + 8·0 g l−1 NH4NO3 + 6·0 g l−1 NaNO318 : 13·38 ± 1·40c0·32 ± 0·01b
24BM + 40 g l−1 CMC + 9·0 g l−1 YE + 2·0 g l−1 NH4NO3 + 9·0 g l−1 NaNO315 : 13·32 ± 1·40c0·22 ± 0·05a
25BM + 40 g l−1 CMC + 12·0 g l−1 YE + 16·0 g l−1 NH4NO3  + 12·0 g l−1 NaNO313 : 12·11 ± 0·50a0·21 ± 0·05a

Effect of initial medium pH

Initial medium pH significantly (< 0·05) affected both BDW and PY (Fig. 6). Penicillium resticulosum Blr1 grew and produced pigment over a broad pH range (3·5–7·5). An initial medium pH outside 5·5–6·5 decreased both BDW and PY. BDW at pH 5·5 (4·42 ± 0·15 g l−1) and 6·5 (3·92 ± 0·31 g l−1) was significantly (< 0·05) higher than that at pH 3·5 (1·38 ± 0·56 g l−1), 4·5 (1·87 ± 0·21 g l−1) or 7·5 (2·30 ± 0·17 g l−1), with no significant (P > 0·05) difference was observed between pH 5·5 and 6·5. A similar trend was observed for PY, which was also significantly (< 0·05) higher at pH 5·5 (0·91 ± 0·12 g l−1) and 6·5 (0·72 ± 0·17 g l−1) than that at pH 3·5 (0·27 ± 0·05 g l−1), 4·5 (0·30 ± 0·05 g l−1) or 7·5 (0·48 ± 0·05 g l−1), but not significantly (P > 0·05) different between pH 5·5 and 6·5.

Figure 6.

Effect of initial pH medium on Penicillium resticulosum Blr1 biomass dry weight and pigment yield (PY) in carboxymethylcellulose–yeast extract–NH4NO3–NaNO3-supplemented corncob hydrolysate culture medium. Values and error bars represent means ± SD (= 5). anova was followed by Tukey's test. a, b, c < 0·05 and *, **, ***, *#, # < 0·05 within respective groups. image, PY; image, biomass.

Effect of incubation period

Incubation period significantly (< 0·05) affected both BDW and PY (Fig 7). BDW and PY increased between 6 and 12 days, then progressively decreased from 12- to 21-days total incubation. BDW at 6 days (1·54 ± 0·29 g l−1) was significantly (< 0·05) lower than that at 9 days (2·46 ± 0·38 g l−1), with both 6- and 9-days BDW significantly (< 0·05) lower than that at 12 (5·17 ± 0·25 g l−1), 15 (5·02 ± 0·59 g l−1), 18 (4·63 ± 0·35 g l−1) and 21 days (5·04 ± 0·85 g l−1). Differences in BDW between 12 and 21 days incubation were insignificant (< 0·05). PY at 6 days (0·26 ± 0·03 g l−1) was also significantly (< 0·05) lower than 9 days (2·46 ± 0·38 g l−1), with both 6 and 9 days yields significantly (< 0·05) lower than 12 (0·58 ± 0·04 g l−1), 15 (0·60 ± 0·04 g l−1), 18 (0·60 ± 0·07 g l−1) or 21 days (0·60 ± 0·01 g l−1), but no significant (< 0·05) difference between 12 and 21 days yields.

Figure 7.

Effect of incubation period on Penicillium resticulosum Blr1 biomass dry weight and pigment yield (PY) in carboxymethylcellulose–yeast extract–NH4NO3–NaNO3-supplemented corncob hydrolysate culture medium. Values and error bars represent means ± SD (n = 3). anova was followed by Tukey's test. a, b, c < 0·05; and *, **, ***< 0·05 within respective groups. Error bars indicate standard deviations. ND, no detection; image, PY; image, biomass.


Filamentous fungi may be identified to species using morphological characteristics (Li et al. 2007). A red pigment producing fungus isolated from Baluran National Park soil was identified as P. resticulosum using such classical taxonomic identification and classification criteria and designated strain Blr1. Dark incubation was required for red pigment production on MEA or CYA, and pigment produced on PDA appeared excreted and diffusible (Fig. 1a). Penicillium sp. are known to exhibit variable responses to light exposure. Szczepanowska and Lovett (1992) found Penicillium notatum yellow-green pigment secretion decreased under dark vs light incubation conditions on Sabouraud dextrose agar. In contrast, Velmurugan et al. (2010) reported dark incubation increased biomass, as well as extracellular and intracellular yellow-red pigment production by Penicillium purpurogenum in a synthetic medium. The apparent production of excretable-diffusible pigment, as observed when P. resticulosum Blr1 was grown on PDA, suggests potential for continuous culture pigment production and harvest from spent medium.

Pigment toxicity and stability are important qualities for wide application in the food industry (Delgado-Vargas et al. 2000; Dufossé 2006). Toxicity assessment is particularly important for fungal-derived pigments, as fungi may produce a wide range of potent toxins. Using the P. resticulosum Blr1 pigment isolated in this study, abnormal clinical symptoms, liver or kidney damage, were not observed in mice fed a ≤500 mg kg−1 pigment dose (in CMC solution) for 28 days, indicating low toxicity (Sopandi and Wardah 2012). In the present study, the red pigment of P. resticulosum Blr1 was stable over a pH range of 5–9, with a progressive 3–11% reduction with lower pH at or below 4. Many investigators have reported pH-stable red pigments from Penicillium. Hailei et al. (2011) similarly reported a red pigment from Penicillium sp. HSD07B stable between pH 2 and 10.

Glucose is generally an excellent carbon substrate for fungal growth, but has been shown to interfere with the biosynthesis of many secondary metabolites (Demain 1986). We found P. resticulusum Blr1 biomass yield approximately 60% higher when cultivated in corncob hydrolysate medium with xylose (59·6%) or CMC (56·7%) as a carbon source, when compared with glucose. Similarly, PY was approximately 150% higher with xylose (161·90%) or CMC (147·62%) compared to glucose. These results are similar to those of Ueno and Ishikawa (1969) who reported higher mycelial weight and pigment percentage from Penicillium islandicum Sopp. grown in Czapek liquid medium with d-xylose as carbon source, compared to d-glucose. Due to increased PY with xylose over glucose as a carbon source, we hypothesized an inhibitory effect of furfural and hydroxymethylfurfural, both by-products of corncob acid hydrolysis, on glucose uptake, biomass production and pigment synthesis. Jeevan et al. (2011) found xylose (71·2–60·5 g l−1) a major product of 2–4% sulphuric acid corncob hydrolysis, as well as glucose (5·1–5·8 g l−1), 5-hydroxymethylfurfural (0·13–0·20 g l−1) and furfural (0·12–0·27 g l−1). Furfural inhibits central enzymes in glycolysis, for example, hexokinase and phosphofructokinase (Banerjee et al. 1981). We found furfural addition to corncob hydrolysate glucose medium progressively decreased glucose consumption (24–68% between 1 and 5 g l−1, respectively), as well as biomass and PYs with increasing furfural concentration (Fig. 4). These results indicate P. resticulosum Blr1 can metabolize xylose via the pentose phosphate pathway in corncob hydrolysate culture medium for biomass and pigment formation. Our observation of greater P. resticulosum Blr1 pigment and biomass production with CMC over glucose as a C source is consistent with Shahriarinour et al. (2011) and Adeleke et al. (2012); who found increased pigment production and growth of cellulolytic fungi on CMC vs glucose containing medium improved screening and isolation. However, Hailei et al. (2011) used a glucose-supplemented potato dextrose broth (PDB) medium with Penicillium sp. HSD07B/C. tropicalis obligate coculture to achieve high (2·75–7·7 g l−1) PYs.

The nitrogen source in growth media plays an important role in microbial growth and enzyme production (Mrudula and Anitharaj 2011). We found P. resticulosum Blr1 had similar biomass and pigment production utilizing organic (peptone and yeast extract) or inorganic (NaNO3, NH4NO3) nitrogen sources. A variety of Penicillium sp. have the reported ability to utilize inorganic and/or organic nitrogen source for pigment production. Gunasekaran and Poorniammal (2008) found peptone the optimal nitrogen source for Penicillium spp. pigment and biomass production. Tariq and Reyaz (2012) found Penicillium chrysogenum pectinase activity greater with an (NH4)2SO4 nitrogen source than peptone and yeast extract. Shivam et al. (2009) found a combination of organic and inorganic nitrogen sources resulted in higher α-galactosidase production compared to either organic or inorganic N sources alone.

Optimal carbon source and C : N ratio for growth vary among fungal species and isolates and may be strain dependent (Gao et al. 2007). We observed maximum biomass (4·55 ± 0·61 g l−1) and PY (0·50 ± 0·06 g l−1) at a C : N ratio of 21 : 1. Wong et al. (1981) reported increased C : N ratio could improve pigment biosynthesis, with Shah et al. (2005) achieving maximum biomass yield in a 35 : 1 C : N medium; while Jackson and Schisler (1992) observed maximum spore yield using a 30 : 1 C : N ratio. Phoolphundh et al. (2010) found N-limited growth conditions significantly enhanced rates of glucose uptake and pigment production by Monascus spp.

Several investigators have found different fungal mycelial morphology during growth in different initial pH media is a critical factor affecting aggregation as well as biomass and pigment formation (Chen and Johns 1993; Duarte and Archer 2003; Unagul et al. 2005; Mendentsev et al. 2005; Boonyapranai et al. 2008). We found maximum P. resticulosum Blr1 biomass and PY in a mild acid (pH 5·5–6·5) medium, within the 5·5–6·0 pH range generally found optimal for growth and pigment production by Penicillium spp. (Mapari et al. 2009), as well as similar to the optimum pH 5·6–6·2 medium for Arpink Red pigment production by P. oxalicum var. Armeniaca CCM 8242 (Dufossé 2006).

Pigments are secondary metabolites generally produced at the end of logarithmic growth or in stationary phase (Bu'Lock 1961; Yongsmith et al. 1999). However, we found pigment formation by P. resticulosum Blr1 started at early logarithmic phase (6 days) and continued until stationary phase (12 days). As pigment or secondary metabolite production has previously been found more closely associated with sporulation (or conidiation) than growth (Stone and Williams 1992; Suhr et al. 2002), we hypothesize conidia formation occurred during logarithmic phase growth, with P. resticulosum Blr1 undergoing microcycle conidiation: the production of conidia directly by spora without intervening hyphal growth (Ulloa and Hanlin 2000), or the immediate recapitulation of sporogenesis after spora germination (Smith et al. 1977). The phenomenon has been described in a number of submerged cultures of asexual fungi (i.e. Aspergillus niger, Paecilomyces uarioti, Penicillium urticae, Neurospora crassa, Penicillium digitatum) (Anderson and Smith 1971; Sekiguchi et al. 1975; Anderson et al. 1978). Induction of microcycle conidiation comprises inhibition of apical growth of germ tube(s) and extension of the conidial swelling stage (mainly via hydration and nutrient uptake) by so-called spherical growth, which is accompanied by active biomass growth, and after its termination, a conidiophore is formed instead of a germ tube (Duncan et al. 1978). Penicillium sp. conidiation involves the apical compartment differentiation into a specialized reproductive cell (phialide), which undergoes successive mitotic divisions, each resulting in a new specialized daughter cell: the conidium (Roncal and Ugalde 2003). Exposure of hyphae to air has been recognized as the most powerful stimulus for conidiogenesis onset in filamentous fungi, including Penicillium sp. (Morton 1961). However, we did not observe increases in floating biomass or other indications of increased exposure of hyphae to air during log phase growth. We found 12-days incubation optimum for P. resticulosum Blr1 pigment production in corncob hydrolysate CMC–yeast extract–NH4NO3–NaNO3 medium. Incubation period for growth and pigment production varies among fungal species or strain and fermentation condition dependent. Dhale and Vijay-Raj (2009) utilized a 10-days incubation time to obtain pigment from Penicillium sp NIOM-02 in malt extract medium at room temperature with 118 rev min−1 agitation, while Jiang et al. (2005) used an incubation time of 14 days (25°C, 180 rev min−1) to obtain red pigment from Penicillium spp.

Penicillium species or strain, medium composition, temperature and incubation time have all been found to influence production of red pigments (Mapari et al. 2006). In the present study, cultivation of P. resticulosum Blr1 in corncob hydrolysate culture medium supplemented with 5·0 mg l−1 KH2PO4, 0·02 g l−1 MgSO4·7H2O, 30 g l−1 CMC or xylose, 3·0 g l−1 yeast extract, 4·0 g l−1 NH4NO3 and 3·0 g l−1 NaNO3 at an initial pH of 5·5 for 12 days at 60–70% r.h. produced a PY of 0·58 ± 0·04 g l−1. Dufossé (2006) achieved 1·5–2·0 g l−1 pigment using P. oxalicum var. Armeniaca CCM 8242 in liquid carbohydrate-based medium (sucrose, molasses, corn extract) with a yeast autolysate or extract nitrogen source, ZnSO4 and MgSO4, pH 5·6–6·2, 27–29°C and 3–4 days incubation. Gunasekaran and Poorniammal (2008) obtained 1·55 g l−1 red PY using Penicillium spp. in soluble starch–peptone liquid medium, while Hailei et al. (2011) achieved 2·75 and 7·7 g l−1 of a pH-stable, nontoxic or mutagenic red pigment in shake flask and 15-l bioreactor culture, respectively, using Penicillium sp. HSD07B/C. tropicalis (obligate) coculture in a glucose-supplemented PDB-based medium. Maximum PY in the present study was approximately 70–90% lower than that from the above studies.

Further structural, physicochemical and toxicological characterization of P. resticulosum Blr1 red pigment chemistry, as well as phylogenetic analysis of the isolate, was outside the present study scope. However, molecular phylogenetic and taxonomic analysis of the isolate, as well as comparison of key characteristics for industrial applicability between the various fungi-produced pigments, such as molar extinction coefficient (ε), solubility and resistance to oxidoreduction, is warranted. Fungal pigment production using other agricultural lignocellulosic waste stream materials such as straw and rice husk as a (hydrolysed) basal medium also warrants study.


Penicillium resticulosum Blr1 demonstrated potential as a microbial producer of a pH-stable, nonacutely toxic red pigment in a corncob waste stream cellulose culture medium at 21 : 1 C : N ratio, pH 5·5–6·0, with yeast extract/NH4NO3/NaNO3 and xylose or CMC supplementation and 12-days dark incubation at a yield of approximately 0·5 g l−1. Furfural, type of carbon source, C and N concentration, initial pH and incubation period can each influence biomass and PY using this organism with C and N source addition to a 21 C : N ratio and initial medium pH 5·5 increasing pigment production. The use of this low-cost agricultural waste stream material as a fermentation substrate base for pigment production represents a potential environmentally sustainable, less temporally susceptible process for agricultural waste reuse while producing a commodity in increasing demand.


The authors thank the Directorate General of Higher Education, National Education Department of Indonesia for funding support through its fundamental research competition. We also thank Mrs. Sue Gill, Mr. Mark Stinson and microbiology section staff, Queensland University of Technology for logistical assistance.