analysis of FTIR-HTS spectra of *Mucor circinelloides* biomass grown under Pi4 level. The loadings of spectral PCA (A), the score plot (B).

**Author Contributions:** Conceived the research idea, V.S., B.Z., and A.K.; Designed the experiments, B.Z., S.D., V.S., and A.K.; methodology, V.S., B.Z., and A.K.; performed the experimental work, S.D.; analyzed the data, V.T. and S.D.; discussed the results, S.D., V.S., B.Z., A.K., and V.T.; wrote the manuscript, S.D.; discussed and revised the manuscript, S.D., V.S., B.Z., V.T., S.J.H., and A.K. All authors have read and agreed to the published version of the manuscript.

**Funding:** This research was funded by Research Council of Norway—FMETEKN Grant, project number 257622; BIONÆR Grant, project numbers 268305, 305215; DAAD Grant, project number 309220; HAVBRUK2 Grant, project number 302543/E40; MATFONDAVTALE Grant, project number 301834/E50.

**Conflicts of Interest:** The authors declare no conflict of interest. The funders had no role in the design of the study; in the collection, analyses, or interpretation of data; in the writing of the manuscript; or in the decision to publish the results.

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Article

# Biotransformation of Animal Fat-By Products into ARA-Enriched Fermented Bioproducts by Solid-State Fermentation of *Mortierella alpina*

Ondrej Slaný¹, Tatiana Klempová ¹,* ID, Volha Shapaval ², Boris Zimmermann ², Achim Kohler ²
and Milan Čertík ¹ID

¹ Institute of Biotechnology, Faculty of Chemical and Food Technology, Slovak University of Technology, Radlinského 9, 812 37 Bratislava, Slovakia; ondrej.slany@stuba.sk (O.S.); milan.certik@stuba.sk (M.Č.)

² Faculty of Science and Technology, Norwegian University of Life Sciences, Postbox 5003, 1432 Ås, Norway; volha.shapaval@nmbu.no (V.S.); boris.zimmermann@nmbu.no (B.Z.); achim.kohler@nmbu.no (A.K.)

* Correspondence: tatiana.klempova@stuba.sk; Tel.: +421-949-582-157

Received: 25 September 2020; Accepted: 17 October 2020; Published: 21 October 2020

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**Abstract:** Solid-state fermentation (SSF) is a powerful fermentation technology for valorizing rest materials and by-products of different origin. Oleaginous *Zygomycetes* fungi are often used in SSF as an effective cell factory able to valorize a wide range of hydrophilic and hydrophobic substrates and produce lipid-enriched bioproducts. In this study, for the first time, the strain *Mortierella alpina* was used in SSF for the bioconversion of animal fat by-products into high value fermented bioproducts enriched with arachidonic acid (ARA). Two cereals-based matrixes mixed with four different concentrations of animal fat by-product were evaluated for finding optimal conditions of a fat-based SSF. All obtained fermented bioproducts were found to be enriched with ARA. The highest substrate utilization (25.8%) was reached for cornmeal and it was almost double than for the respective wheat bran samples. Similarly, total fatty acid content in a fermented bioproduct prepared on cornmeal is almost four times higher in contrast to wheat bran-based bioproduct. Although in general the addition of an animal fat by-product caused a gradual cessation of ARA yield in the obtained fermented bioproduct, the content of ARA in fungal biomass was higher. Thus, *M. alpina* CCF2861 effectively transformed exogenous fatty acids from animal fat substrate to ARA. Maximum yield of 32.1 mg of ARA/g of bioproduct was reached when using cornmeal mixed with 5% (w/w) of an animal fat by-product as substrate. Furthermore, implementation of attenuated total reflectance Fourier transform infrared (ATR-FTIR) spectroscopy in characterization of obtained SSF bioproducts was successfully tested as an alternative tool for complex analysis, compared to traditional time-consuming methods.

**Keywords:** *Mortierella alpina*; solid-state fermentation; animal fat by-product; arachidonic acid; ATR-FTIR spectroscopy

## 1. Introduction

In Europe, approximately 16 million tonnes of animal fat (AF) by-products are processed annually by meat processors and renderers [1]. The total amount of AF by-products is expected to increase continuously, since consumers and health authorities in European countries focus on highly unsaturated fat diets [2]. In addition, due to the population growth, meat production in Europe is expected to grow. Even though some of these AF by-products comply with EU standards (EC No. 1069/2009) and can be served for human consumption, they are used almost exclusively as ingredients of animal feed. The biggest problem lies in the composition of AF by-products since they are rich in saturated and monounsaturated fatty acids. Long-term consumption of saturated fat elevates the risk of certain
diseases such as diabetes or cardio-vascular problems. Thus, human food, as well as animal and aquaculture feeds, need to be supplied with the essential polyunsaturated ω-3 and ω-6 fatty acids (PUFAs), such as linoleic acid (LA; C18:2, ω-6), α-linolenic acid (ALA; C18:3, ω-3), arachidonic acid (ARA; C20:4, ω-6), eicosapentaenoic acid (EPA, C20:5, ω-3) and docosahexaenoic acid (DHA, C22:6, ω-3) [3]. Therefore, the development of sustainable processes for valorizing AF by-products into high-value bioproducts is urgently needed.

Fungal fermentation is an emerging technology for upgrading different types of by-products [4,5]. A certain group of filamentous fungi can accumulate up to 85% (w/w) of triacylglycerols (TAGs) containing high amount of essential PUFAs as a storage material of the cells [6]. Fungal biomass produced with these oleaginous fungi contains single-cell oils with a high share of essential ω-3 and ω-6 PUFAs, and is considered a high-value bioproduct with applications in food, feed and nutraceutical industry. Fungal biomass can be obtained by two fermentation processes: submerged or solid-state fermentation (SSF).

SSF is a fermentation technology using solid or semi-solid substrates in the absence of any water or using a low level of free-flowing water [7]. SSF has many advantages over more common submerged fermentation: (1) low energy requirements are accompanied by the high product yield; (2) downstream costs are significantly lower due to the high product applicability; (3) SSF conditions are more favorable for the microbial growth as they resemble the natural environment of microorganisms, resulting in a better performance of the fermentation process; (4) due to the high product/volume productivity of SSF, smaller fermentation volumes are possible [8]; (5) consumption of a lower amount of water and generation of low to almost zero amount of waste [9]. Solid substrates used in SSF frequently originate from agro-industrial waste materials and they serve as support materials for the optimal fungal growth, proliferation and single-cell oil production. Commonly used SSF substrates are cereal-based materials, such as barley flakes, millet grain, wheat bran, cornmeal or oat flakes [10,11]; fruit or vegetable-based materials, such as pulp, pomaces and peels (i.e., residues from plums, pineapple, carrot, papaya etc.) [12,13]; or they often come as residue from other types of industry, for example spent malt grain, groundnut fodder, forestry rests or wooden sawdust [7,14]. Several studies investigated the suitability of different oleaginous fungi, belonging to orders *Mucorales* and *Mortierellales*, for converting various waste materials and by-products based substrates by SSF into fermented BPs enriched with a wide range of essential PUFAs, such as γ-linolenic acid (GLA; C18:3, n-6), ARA or EPA [10,12,15–18]. Especially cereal-based solid matrixes are very suitable for these oleaginous fungi due to their chemical composition, such as the presence of easily accessible carbon, organic nitrogen and other macro- and micro-nutrients. Cereal-based matrixes provide a useful source for good fungal proliferation, hyphae penetration and stable lipid accumulation in fungal cells [10,15,16].

Oleaginous fungus *Mortierella alpina* is a well-known and thoroughly described species with a high capacity of lipid accumulation and good ability to produce industrially relevant essential PUFAs, especially ARA in both submerged fermentations and SSF [19–21]. It has also been reported that *M. alpina* is able to incorporate and transform exogenous fatty acids [22]. Thus, in order to improve the conversion of solid wastes into ARA-enriched bioproduct, supplementation with exogenous oils containing precursors of ARA is advised. Thus, the addition of vegetable oils, such as sunflower, rapeseed, corn, soybean or linseed oil containing individual fatty acid precursors of ARA led to a rapid increase of ARA yield in the final fermented bioproduct obtained by fungal SSF [10]. Such knowledge led to the hypothesis that the oleaginous fungus *M. alpina* should also be able to utilize and convert solid lipidic waste, such as AF by-products.

When developing a new SSF process and optimizing SSF processes, it is crucial to monitor key process parameters, such as humidity, airflow, oxygen transfer and controlling the quality of final fermented bioproduct. Quality parameters of the final fermented BP are fatty acid profile, total lipid content, amount of fungal biomass and total biochemical profile. For the determination of fatty acid profile and content, gas chromatography coupled with flame ionization detector (GC-FID) is the most commonly used technique [23], while the total amount of fungal biomass obtained by SSF is frequently
analyzed by the glucosamine method [24]. Fourier transform infrared (FTIR) spectroscopy is a new emerging technique that has already been extensively used for characterizing biomass from different types of submerged cultivations [25–28]. FTIR spectroscopy is a biophysical technique allowing to measure the biomass in its native form. It is non-destructive and rapid and does not require extraction of single components. Therefore, the characterization of a bioproduct obtained in the process of SSF using FTIR spectroscopy could bring the new possibilities in complex monitoring of SSF process.

To the authors knowledge, this is the first study investigating the possibility to develop an efficient bioconversion of AF by-products into functional lipid-rich fermented bioproduct by SSF using *M. alpina*. Furthermore, the presented study introduces, describes and evaluates FTIR spectroscopy as an alternative method for accurate SSF monitoring.

## 2. Materials and Methods

### 2.1. Production Microorganism and Preparation of Spore Suspension

The oleaginous fungal strain *Mortierella alpina* CCF2861 used in this study was obtained from culture collection of fungi (CCF, Charles University, Prague, Czech Republic). The strain was kept on potato-dextrose agar media (PDA, Carl Roth, Germany) at 4 °C and regularly re-inoculated every 3 months.

The spore suspension for the inoculation of the fermentation substrate was prepared from a 14-day-old culture grown on PDA medium. Spores have been washed using an aqueous solution of 0.05% Tween®® 40 and suspension was diluted to achieve a final concentration of 10⁶ spores/mL.

### 2.2. Conditions of Solid-State Fermentation

SSF cultivation was performed in the microporous high-density polyethylene bags (20 × 30 cm) containing 20 g of dry cereal-based matrix, to which various amounts of AF by-products (Norilia, Oslo, Norway) mixed with Tween®® 40 (Sigma-Aldrich, Darmstadt, Germany) were added. SSF experiments were performed using two waste cereal-based matrixes: cornmeal (CM; Amylum Slovakia, Boleráz, Slovakia) and wheat brans (WB; Mill Pohronský Ruskov, Pohronský Ruskov, Slovakia). The composition of AF used is summarized in Table 1.

<table><caption>**Table 1.** Fatty acid (FA) composition and percentage of individual lipid components of animal fat (AF) by-products used for the solid-state fermentation by *Mortierella alpina*.</caption><thead><tr><th>FA</th><th>[%]</th></tr></thead><tbody><tr><td>C14:0</td><td>2.23</td></tr><tr><td>C16:0</td><td>25.66</td></tr><tr><td>C16:1 n-7</td><td>2.18</td></tr><tr><td>C18:0</td><td>21.33</td></tr><tr><td>C18:1 n-9</td><td>38.34</td></tr><tr><td>C18:1 n-7</td><td>1.94</td></tr><tr><td>C18:2 n-6</td><td>5.47</td></tr><tr><td>C18:3 n-3</td><td>0.81</td></tr><tr><td>C20:0</td><td>0.23</td></tr><tr><td>Other fatty acids</td><td>1.81</td></tr><tr><th colspan="2">Lipid Structure</th></tr><tr><td>Polar lipids</td><td>0.14</td></tr><tr><td>Monoacylglycerols</td><td>0.28</td></tr><tr><td>Diacylglycerols</td><td>2.08</td></tr><tr><td>Sterol structures</td><td>9.38</td></tr><tr><td>Free fatty acids</td><td>7.20</td></tr><tr><td>Triacylglycerols</td><td>66.54</td></tr><tr><td>Esterified sterols</td><td>11.23</td></tr><tr><td>Other lipid structures</td><td>3.15</td></tr></tbody></table>
The cereal materials were soaked for 2 h in 20 mL (ratio solid to liquid 1:1) of either distilled water, Tween 40®® solution or AF emulsion, as per Table 2. After 2 h, all SSF substrates in bags were autoclaved (105 °C, 45 min). After cooling down, the substrates were inoculated with 4 mL of spore suspension of *M. alpina*. The fermentation ran for 10 days at 20 °C. In order to maintain the homogenous growth of fungi, substrates were mixed thoroughly after inoculation. During the fermentation process, the whole content of the fermentation bags was gently mixed once a day. All experiments were performed in three independent biological replicates.

<table><caption>**Table 2.** Amount of added Tween®® 40 and animal fat to cereal matrixes used for the solid-state fermentation by *Mortierella alpina*.</caption><thead><tr><th>Cereal Matrix</th><th>Tween®® 40<br>[%(w/w)]</th><th>Animal Fat<br>[%(w/w)]</th></tr></thead><tbody><tr><td rowspan="8">cornmeal</td><td>0</td><td>0<sup>1</sup></td></tr><tr><td>0.5</td><td>0<sup>2</sup></td></tr><tr><td>1</td><td>0<sup>2</sup></td></tr><tr><td>2</td><td>0<sup>2</sup></td></tr><tr><td>3</td><td>0<sup>2</sup></td></tr><tr><td>0.5</td><td>5<sup>3</sup></td></tr><tr><td>1</td><td>10<sup>3</sup></td></tr><tr><td>2</td><td>20<sup>3</sup></td></tr><tr><td>3</td><td>30<sup>3</sup></td></tr><tr><td rowspan="8">wheat bran</td><td>0</td><td>0<sup>1</sup></td></tr><tr><td>0.5</td><td>0<sup>2</sup></td></tr><tr><td>1</td><td>0<sup>2</sup></td></tr><tr><td>2</td><td>0<sup>2</sup></td></tr><tr><td>3</td><td>0<sup>2</sup></td></tr><tr><td>0.5</td><td>5<sup>3</sup></td></tr><tr><td>1</td><td>10<sup>3</sup></td></tr><tr><td>2</td><td>20<sup>3</sup></td></tr><tr><td>3</td><td>30<sup>3</sup></td></tr></tbody></table>

## 2.3. Preparation of AF Emulsion

Due to the fact that cereal-based matrixes have a hydrophilic nature while AF is hydrophobic, there is a need to perform the pretreatment of the fat materials before mixing it with cereal matrixes for obtaining homogenous substrates for SSF. The pretreatment of AF by-product was performed by preparing homogenous fat-water-Tween®® 40 emulsions, where the emulsifier Tween®® 40 formed a film around the dispersed lipid droplets and thereby reduced interfacial tension [29].

AF emulsions were prepared from distilled water, Tween®® 40 and AF using a combination of heating (80 °C, 10 min, stirring) and sonication (15 min) using VWR USC300T sonicator (VWR International, Radnor, PA, USA).

## 2.4. Humidity and Substrate Utilization Analysis

The obtained fermented bioproducts were collected and the humidity was measured. The humidity of substrates was measured by Moisture Analyzer Radwag 50/1. R (Radwag, Radom, Poland). Subsequently, the substrates were dried at 65 °C until a constant weight was achieved. The substrate utilization was calculated from the weight difference of the dry non-fermented substrate (control) and dry fermented bioproducts. Each sample was homogenized using a blender and stored in sterile Falcon tubes at laboratory temperature before the analysis was performed.

<footnote n="1">¹ one-component samples (distilled water); ² two-component samples (distilled water + Tween®® 40);</footnote>

<footnote n="3">³ three-component samples (distilled water + Tween®® 40 + AF).</footnote>
## 2.5. Estimation of Fungal Biomass in Fermented Bioproduct

To estimate the amount of fungal biomass in the fermented bioproducts, the method based on the estimation of glucosamine (GlcN) content was used [30,31]. In the first step, alkali insoluble material (AIM) was prepared according to Zamani et al. [24]: 0.5 M NaOH solution (3 mL) was added to 100 mg of each sample of fermented BP and the mixtures were heated at 90 °C for 16 h. Subsequently, samples were centrifuged (5000 rpm, 10 min) and washed 5 times with distilled water. Supernatants were removed and obtained AIMs were dried for 36 h at 75 °C. Further, AIM samples were hydrolyzed by adding 5 mL of 6M hydrochloric acid and incubating at 100 °C for 12 h. The analysis of glucosamine (GlcN) from the hydrolyzed AIM samples was performed spectrophotometrically according to Aidoo et al. [30] and GlcN content was calculated according to the standard calibration curve. Each analysis was performed in three independent technical replicates.

## 2.6. Microsporic Observation of Fungal Mycelia during SSF

The snapshots of fermented bioproducts were obtained by Dino-Lite Pro USB microscope. Fungal mycelia of *M. alpina* was cut out from the SSF matrix, suspended in 500 µL of sterile distilled water and observed by a confocal microscope Leica DM750 (Leica microsystems Ltd., Wetzlar, Germany), equipped with Leica HI PLAN 100x/1.25 oil objective lens (Leica microsystems Ltd., Wetzlar, Germany). Snapshots of individual hyphae were carried out with a Leica DFC290HD digital camera (Leica microsystems Ltd., Wetzlar, Germany).

## 2.7. Analysis of Fatty Acid Profile Andf Content in Fermented Bioproduct

Fatty acids (FA) from the SSF matrixes were converted into their methyl esters (FAMEs) by a modified method of Čertík and Shimizu [32]: 20 mg of dry homogenized bioproducts were mixed with 1 mL of dichloromethane containing 0.1 mg of heptadecanoic acid as an internal standard and 2 mL of anhydrous methanolic HCl solution. Samples were incubated at 50 °C for 3 h. After cooling down, 1 mL of distilled water was added and FAMEs were extracted with 1 mL of hexane. FAMEs were subsequently analyzed by gas chromatography according to the method described by Gajdoš et al. [33]. The identification of the FAMEs peaks was performed by comparison with authentic standards of C4-C24 FAME mixtures (Sigma Aldrich, USA). Quantitative evaluation of individual and total fatty acids was performed using an internal standard of heptadecanoic acid (C17:0, Sigma-Aldrich, Darmstadt, Germany) and calculated by ChemStation B 01 03 (Agilent Technologies, Santa Clara, CA, USA). Each analysis was performed in three independent technical replicates.

## 2.8. Lipid Isolation and Analysis of Lipid Classes by TLC

The lipids for the analysis of lipid classes were extracted with chloroform/methanol (2:1, *v*/*v*) according to original Folch et al. method [34] modified by Klempová et al. [15]. The lipid extracts were loaded on TLC silica plates 60 (Merck, Germany) using CAMAG TLC Sampler 4 (CAMAG, Muttenz, Switzerland) and TLC analysis was performed according to the method described by Gajdoš et al. [33]. Separated lipid fractions were analyzed densitometrically using CAMAG TLC Scanner 4 (CAMAG, Muttenz, Switzerland) and quantified using WinCATS software (CAMAG, Muttenz, Switzerland). Each analysis was performed in three independent technical replicates.

## 2.9. ATR-FTIR Spectroscopy Analysis

Fourier transform infrared (FTIR) spectroscopy was done employing an attenuated total reflectance (ATR) accessory for profiling the total biochemical composition of the obtained fermented bioproducts. ATR-FTIR measurements were performed using a Vertex 70 FTIR spectrometer (Bruker Optic, Billerica, MA, USA) with a single-reflection attenuated total reflectance (SR-ATR) accessory. The ATR-FTIR spectra were recorded with 32 scans using a horizontal SR-ATR diamond prism with 45° angle of incidence on a Specac (Slough, UK) High Temperature Golden Gate ATR Mk II. Of each homogenized
dried fermented sample and of each non-fermented substrate sample, 10 mg of sample mass was prepared by homogenization with 2.8 mm stainless steel grinding balls (OPS Diagnostics, USA) using tissue homogenizer (5800 rpm, cycle 2 × 15 s with a 30 s pause) Percellys Evolution (Bertin Instruments, France). The samples were subsequently transferred to the surface of the ATR crystal. All ATR-FTIR measurements were performed in five technical replicates, resulting in 360 spectra in total. Spectra were recorded in a region between 7000 and 600 cm⁻¹ with a resolution of 4 cm⁻¹. Each spectrum was recorded as the ratio of the sample spectrum to the spectrum of the empty ATR plate. The recording of spectra was performed using the OPUS 7.5 software (Bruker Optic, Billerica, MA, USA).

## 2.10. Data Analysis

The fatty acid GC-FID and GlcN data were analysed by ANOVA using Microsoft Excel (Microsoft Office 365 software pack) equipped with a data analysis tool. Post-hoc testing was performed for the ANOVA results using Tukey's HSD test in programming language R and in Python v. 3.7 using StatsModels libraries.

FTIR-ATR spectra were obtained for the region of 4000–600 cm⁻¹, selected as the spectral region containing bands distinctive for lipids, proteins and polysaccharides. The analysis of FTIR-ATR spectra was performed by first applying pre-processing and then principle component analysis (PCA). The pre-processing was performed by transforming to second-derivative spectra using the Savitzky–Golay algorithm with a polynomial of degree 2 and a window size of 11. The second-derivative spectra were pre-processed by extended multiplicative signal correction (EMSC) [35,36]. PCA was performed for three spectral regions, lipid (3050–2800 cm⁻¹ combined with 1800–1700 cm⁻¹), protein (1700–1500 cm⁻¹) and polysaccharide (1200–700 cm⁻¹), using 11 principle components. The following software packages were used for data analysis: Unscrambler X version 10.5.1 (CAMO Analytics, Norway), Orange data mining toolbox version 3.24 (University of Ljubljana, Slovenia) [37].

# 3. Results

The present study is focused on utilization of AF by-products by the process of SSF employing fungus *M. alpina*. We evaluated two single-component control substrates, eight two-component control substrates containing Tween®® 40 and eight three-component animal fat-based substrate mixtures (Table 1). Since emulsifier Tween®® 40 may have impacted the fungal metabolism, the effect of Tween®® 40 itself was also investigated.

## 3.1. Fungal Growth and Aubstrate Utilization during SSF

It was observed that the presence of Tween 40 at different concentrations in cereal-based substrates had no significant impact on the substrate utilization, substrate humidity, pH, biomass growth and FA content (*p*-value > 0.4; F parameter > Fcrit) (Table 2) and productivity was comparable with the control substrates based on cereal matrixes only.

It was also found that the utilization of control substrates and AF-based substrates in SSF varied depending on the type of cereal matrix used and the amount of AF added (Table 3). Generally, higher substrate utilization and fungal biomass growth was observed for the single-component and two-component substrates. The highest substrate utilization (25.8%) was reached for the substrates containing cornmeal (CM) and 1% of Tween (Table 3) and the highest fungal growth (289.3 mg/g of bioproducts) was obtained for the substrate containing CM with 3% of Tween. The utilization of CM single- and two-component control substrates was almost twice higher than for the respective wheat bran (WB) substrates (Table 3). Similarly, the fungal growth was 1.5-times higher using CM comparing to WB.
<table><caption>**Table 3.** Substrate utilization, substrate humidity, pH and fungal biomass (FBM) content in fermented bioproducts obtained from SSF of animal fat (AF) by-product by *Mortierella alpina*.</caption><thead><tr><th></th><th>Substrate Utilization [%]</th><th>Substrate Humidity [%]</th><th>pH</th><th>FBM [mg/g BP]</th></tr></thead><tbody><tr><td>cornmeal (CM)</td><td>22.3 ± 0.9</td><td>60.0 ± 0.5</td><td>5.4 ± 0.1</td><td>241.3 ± 10.5</td></tr><tr><td>CM + 0.5% Tween 40</td><td>23.1 ± 0.5</td><td>58.1 ± 0.6</td><td>5.5 ± 0.0</td><td>244.0 ± 26.8</td></tr><tr><td>CM + 1% Tween 40</td><td>25.8 ± 0.5</td><td>60.1 ± 0.7</td><td>5.3 ± 0.1</td><td>272.8 ± 13.5</td></tr><tr><td>CM + 2% Tween 40</td><td>24.3 ± 0.6</td><td>59.0 ± 1.5</td><td>5.3 ± 0.3</td><td>286.2 ± 6.1</td></tr><tr><td>CM + 3% Tween 40</td><td>23.7 ± 1.0</td><td>58.6 ± 0.8</td><td>5.2 ± 0.1</td><td>289.3 ± 13.8</td></tr><tr><td>CM + 0.5% Tween 40 + 5% AF</td><td>21.1 ± 1.4</td><td>53.2 ± 0.6</td><td>5.2 ± 0.0</td><td>137.1 ± 15.6</td></tr><tr><td>CM + 1% Tween 40 + 10% AF</td><td>17.6 ± 0.4</td><td>48.8 ± 1.3</td><td>4.6 ± 0.1</td><td>120.7 ± 10.3</td></tr><tr><td>CM + 2% Tween 40 + 20% AF</td><td>13.1 ± 0.4</td><td>40.0 ± 1.1</td><td>4.5 ± 0.2</td><td>82.7 ± 14.8</td></tr><tr><td>CM + 3% Tween 40 + 30% AF</td><td>10.7 ± 0.4</td><td>36.5 ± 1.5</td><td>4.3 ± 0.1</td><td>73.7 ± 6.0</td></tr><tr><td>wheat bran (WB)</td><td>13.2 ± 0.5</td><td>58.7 ± 1.9</td><td>5.5 ± 0.1</td><td>188.8 ± 1.4</td></tr><tr><td>WB + 0.5% Tween 40</td><td>12.3 ± 0.6</td><td>58.9 ± 0.1</td><td>5.1 ± 0.5</td><td>174.2 ± 10.4</td></tr><tr><td>WB + 1% Tween 40</td><td>12.5 ± 0.4</td><td>56.8 ± 1.9</td><td>5.3 ± 0.2</td><td>193.6 ± 5.2</td></tr><tr><td>WB + 2% Tween 40</td><td>12.6 ± 0.7</td><td>58.4 ± 0.2</td><td>5.2 ± 0.2</td><td>168.1 ± 5.9</td></tr><tr><td>WB + 3% Tween 40</td><td>13.6 ± 0.6</td><td>54.2 ± 4.5</td><td>5.4 ± 0.3</td><td>166.0 ± 1.1</td></tr><tr><td>WB + 0.5% Tween 40 + 5% AF</td><td>10.4 ± 0.4</td><td>54.4 ± 1.0</td><td>5.1 ± 0.2</td><td>205.0 ± 1.1</td></tr><tr><td>WB + 1% Tween 40 + 10% AF</td><td>9.6 ± 0.2</td><td>48.7 ± 1.6</td><td>4.8 ± 0.1</td><td>189.9 ± 4.8</td></tr><tr><td>WB + 2% Tween 40 + 20% AF</td><td>9.3 ± 0.7</td><td>44.2 ± 2.2</td><td>4.5 ± 0.1</td><td>122.7 ± 5.1</td></tr><tr><td>WB + 3% Tween 40 + 30% AF</td><td>8.8 ± 0.5</td><td>39.1 ± 0.7</td><td>4.3 ± 0.0</td><td>92.2 ± 5.2</td></tr></tbody></table>

## 3.2. Microscopic Observation of SSF Process

Fungal growth during SSF was also observed microscopically (Figure 1a,b). Similar morphology characteristics of *M. alpina* grown on control single- and two-component substrates and AF supplemented substrates were found. *M. alpina* was able to completely cover the substrate surface and its hyphae also sufficiently penetrated inside the substrate. Moreover, the presence of lipid structures formed in individual fungal hyphae was detected (Figure 1c).

<figure><img src="image_86.png" /><figcaption>**Figure 1.** Microscopic images of *M. alpina* hyphae covering wheat bran (a), wheat bran with animal fat during solid-state fermentation (b) and lipid droplets accumulated inside *M. alpina* hyphae (c).</figcaption></figure>
### 3.3. *The Impact of AF Supplementation on the Humidity and pH of the Fermented Bioproducts*

It was found that increasing the amount of AF in substrate mixtures reduced the humidity of the fermented bioproducts (Table 3). Fermented bioproducts obtained from the SSF of control substrates had pH 5.3 (CM) and pH 5.5 (WB), while fermented bioproducts obtained from SSF of AF-based substrates showed lower pH values. The increase in AF amounts in fermentation substrates strongly affected the pH values of fermented bioproducts and led to the gradual reduction of pH in the SSF system (Table 3). The lowest pH (4.3) was observed for both cereal-based fermented bioproducts with a 30% (w/w) AF supplementation.

### 3.4. *Lipid Profile and Accumulation*

We observed that for the production of fatty acid-rich fungal biomass the CM is a more suitable substrate compared to WB (Table 3). Although the total fatty acid content in both substrates was similar, the total fatty acids content in bioproducts derived from SSF of single and two-component control substrates based on CM was almost four times higher in contrast to WB-based bioproducts. The total fatty acid content in fermented WB-based bioproducts (single- and two-component) was always lower after fermentation (Table 4). In all three-component bioproducts, the accumulation of fatty acids was strongly affected by the presence of AF, since the added AF was homogenized with other substrate components (Table 4). In addition, it was observed that the total fatty acid content of fermented bioproducts still varied depending on the cereal matrix used. When CM was used as a matrix, the total fatty acid content in fermented bioproducts was elevated compared to the non-fermented substrate. When WB was used as a matrix, the total fatty acid content dropped in fermented bioproducts comparing to the non-fermented substrate. Nevertheless, these results suggest that fungal strain *M. alpina* CCF2861 is definitively able to utilize exogenous fatty acids of AF by-product added to the substrate not only for its own fatty acids biosynthesis, but for hyphae proliferation and growth as well.
<table><caption>**Table 4.** Total fatty acid (TFA) content and fatty acid composition of non-fermented (nf) and fermented (f) cereal substrates (cornmeal—CM, wheat bran—WB) with the addition of different amounts of Tween 40 and animal fat (AF). The results are average of three independent biological replicates with α < 5%.</caption><thead><tr><th rowspan="2"></th><th colspan="2">TFA</th><th colspan="10">Fatty Acids (%)</th></tr><tr><th>(%/BP)</th><th>C14:0</th><th>C16:0</th><th>C16:1, n-7</th><th>C18:0</th><th>C18:1, n-9</th><th>C18:1, n-7</th><th>C18:2, n-6</th><th>C18:3, n-6</th><th>C18:3, n-3</th><th>C20:3, n-6</th><th>C20:4, n-6</th><th>Others</th></tr></thead><tbody><tr><td colspan="14"><strong>cornmeal</strong></td></tr><tr><td>CM nf</td><td>3.2</td><td>0.6</td><td>12.7</td><td>nd</td><td>2.5</td><td>27.3</td><td>0.7</td><td>54.7</td><td>nd</td><td>1.5</td><td>nd</td><td>nd</td><td>nd</td></tr><tr><td>CM f</td><td>13.8</td><td>0.8</td><td>14.5</td><td>0.1</td><td>5.6</td><td>13.7</td><td>0.7</td><td>23.8</td><td>2.3</td><td>0.5</td><td>2.5</td><td>26.4</td><td>9.0</td></tr><tr><td>+0.5%Tween 40 nf</td><td>3.7</td><td>0.7</td><td>14.3</td><td>nd</td><td>2.4</td><td>26.2</td><td>0.7</td><td>54.1</td><td>nd</td><td>1.5</td><td>nd</td><td>nd</td><td>nd</td></tr><tr><td>+0.5%Tween 40 f</td><td>13.7</td><td>0.6</td><td>15.6</td><td>0.1</td><td>5.8</td><td>13.2</td><td>1.0</td><td>23.5</td><td>2.4</td><td>0.3</td><td>2.5</td><td>27.5</td><td>7.6</td></tr><tr><td>+1%Tween 40 nf</td><td>3.4</td><td>0.8</td><td>16.6</td><td>nd</td><td>2.6</td><td>25.4</td><td>0.7</td><td>52.4</td><td>nd</td><td>1.6</td><td>nd</td><td>nd</td><td>nd</td></tr><tr><td>+1%Tween 40 f</td><td>13.4</td><td>0.8</td><td>16.0</td><td>0.1</td><td>5.7</td><td>12.8</td><td>0.6</td><td>22.6</td><td>2.3</td><td>0.4</td><td>2.6</td><td>27.1</td><td>8.9</td></tr><tr><td>+2%Tween 40 nf</td><td>4.3</td><td>1.0</td><td>20.3</td><td>nd</td><td>2.6</td><td>23.7</td><td>0.6</td><td>50.2</td><td>nd</td><td>1.5</td><td>nd</td><td>nd</td><td>nd</td></tr><tr><td>+2%Tween 40 f</td><td>13.8</td><td>0.8</td><td>17.3</td><td>0.1</td><td>5.8</td><td>13.0</td><td>0.6</td><td>22.7</td><td>2.3</td><td>0.5</td><td>2.5</td><td>26.1</td><td>8.4</td></tr><tr><td>+3%Tween 40 nf</td><td>4.7</td><td>0.6</td><td>23.9</td><td>nd</td><td>2.8</td><td>22.3</td><td>0.6</td><td>48.3</td><td>nd</td><td>1.3</td><td>nd</td><td>nd</td><td>nd</td></tr><tr><td>+3%Tween 40 f</td><td>13.7</td><td>0.8</td><td>18.2</td><td>0.1</td><td>5.7</td><td>12.7</td><td>0.6</td><td>23.0</td><td>2.3</td><td>0.5</td><td>2.5</td><td>25.4</td><td>8.2</td></tr><tr><td>+0.5%T + 5%AF nf</td><td>7.1</td><td>0.7</td><td>18.0</td><td>0.8</td><td>7.8</td><td>30.9</td><td>1.1</td><td>39.4</td><td>nd</td><td>1.3</td><td>nd</td><td>nd</td><td>nd</td></tr><tr><td>+0.5%T + 5%AF f</td><td>14.2</td><td>1.0</td><td>17.1</td><td>0.6</td><td>8.8</td><td>18.0</td><td>1.2</td><td>18.6</td><td>2.1</td><td>0.6</td><td>1.8</td><td>22.6</td><td>7.5</td></tr><tr><td>+1%T + 10%AF nf</td><td>11.3</td><td>1.2</td><td>22.2</td><td>1.7</td><td>12.0</td><td>33.4</td><td>1.5</td><td>26.9</td><td>nd</td><td>1.2</td><td>nd</td><td>nd</td><td>nd</td></tr><tr><td>+1%T + 10%AF f</td><td>16.1</td><td>1.1</td><td>18.8</td><td>1.0</td><td>11.3</td><td>21.1</td><td>1.5</td><td>16.4</td><td>2.0</td><td>0.6</td><td>1.4</td><td>18.3</td><td>6.4</td></tr><tr><td>+2%T + 20%AF nf</td><td>18.1</td><td>1.7</td><td>24.7</td><td>2.0</td><td>15.0</td><td>36.4</td><td>1.7</td><td>17.5</td><td>nd</td><td>1.1</td><td>nd</td><td>nd</td><td>nd</td></tr><tr><td>+2%T + 20%AF f</td><td>19.0</td><td>1.4</td><td>21.0</td><td>1.3</td><td>13.5</td><td>24.8</td><td>1.7</td><td>12.9</td><td>1.6</td><td>0.7</td><td>1.1</td><td>14.6</td><td>5.3</td></tr><tr><td>+3%T + 30%AF nf</td><td>24.3</td><td>1.9</td><td>25.8</td><td>2.1</td><td>16.6</td><td>37.4</td><td>1.8</td><td>13.3</td><td>nd</td><td>1.0</td><td>nd</td><td>nd</td><td>nd</td></tr><tr><td>+3%T + 30%AF f</td><td>24.7</td><td>1.6</td><td>22.4</td><td>1.6</td><td>14.7</td><td>30.4</td><td>1.7</td><td>12.7</td><td>1.2</td><td>0.8</td><td>0.7</td><td>8.6</td><td>3.6</td></tr><tr><td colspan="14"><strong>wheat bran</strong></td></tr><tr><td>WB nf</td><td>3.5</td><td>1.7</td><td>17.1</td><td>nd</td><td>1.0</td><td>17.3</td><td>1.5</td><td>56.1</td><td>nd</td><td>4.5</td><td>nd</td><td>nd</td><td>0.7</td></tr><tr><td>WB f</td><td>3.1</td><td>0.2</td><td>12.6</td><td>0.1</td><td>1.6</td><td>15.3</td><td>1.1</td><td>38.6</td><td>2.2</td><td>2.3</td><td>0.6</td><td>20.4</td><td>5.2</td></tr><tr><td>+0.5%Tween 40 nf</td><td>3.6</td><td>nd</td><td>20.7</td><td>nd</td><td>1.1</td><td>16.6</td><td>1.5</td><td>54.7</td><td>nd</td><td>4.4</td><td>nd</td><td>nd</td><td>0.9</td></tr><tr><td>+0.5%Tween 40 f</td><td>3.4</td><td>0.1</td><td>13.9</td><td>nd</td><td>1.5</td><td>15.9</td><td>1.1</td><td>38.2</td><td>1.8</td><td>2.6</td><td>0.3</td><td>20.5</td><td>4.1</td></tr><tr><td>+1%Tween 40 nf</td><td>3.8</td><td>nd</td><td>21.8</td><td>0.2</td><td>1.2</td><td>16.7</td><td>1.4</td><td>53.4</td><td>nd</td><td>4.2</td><td>nd</td><td>nd</td><td>1.2</td></tr><tr><td>+1%Tween 40 f</td><td>3.3</td><td>0.3</td><td>15.7</td><td>nd</td><td>1.7</td><td>14.4</td><td>1.1</td><td>36.8</td><td>2.3</td><td>2.2</td><td>0.6</td><td>20.6</td><td>4.3</td></tr><tr><td>+2%Tween 40 nf</td><td>3.8</td><td>nd</td><td>26.7</td><td>nd</td><td>1.5</td><td>15.8</td><td>1.4</td><td>50.1</td><td>nd</td><td>3.9</td><td>nd</td><td>nd</td><td>0.7</td></tr><tr><td>+2%Tween 40 f</td><td>3.6</td><td>0.3</td><td>15.5</td><td>nd</td><td>2.1</td><td>13.8</td><td>1.1</td><td>34.9</td><td>2.4</td><td>2.1</td><td>0.2</td><td>22.3</td><td>5.5</td></tr><tr><td>+3%Tween 40 nf</td><td>4.6</td><td>0.2</td><td>33.6</td><td>0.2</td><td>1.8</td><td>13.7</td><td>1.2</td><td>44.9</td><td>nd</td><td>3.5</td><td>nd</td><td>nd</td><td>1.0</td></tr><tr><td>+3%Tween 40 f</td><td>4.0</td><td>0.3</td><td>18.8</td><td>0.2</td><td>2.3</td><td>13.0</td><td>1.0</td><td>31.5</td><td>2.1</td><td>2.0</td><td>0.7</td><td>23.1</td><td>5.1</td></tr><tr><td>+0.5%T + 5%AF nf</td><td>6.6</td><td>1.0</td><td>22.1</td><td>1.0</td><td>9.5</td><td>26.5</td><td>1.6</td><td>34.6</td><td>nd</td><td>2.9</td><td>nd</td><td>nd</td><td>0.8</td></tr><tr><td>+0.5%T + 5%AF f</td><td>5.6</td><td>0.5</td><td>17.2</td><td>0.7</td><td>7.1</td><td>18.2</td><td>1.9</td><td>21.8</td><td>3.5</td><td>1.2</td><td>0.8</td><td>19.6</td><td>7.4</td></tr><tr><td>+1%T + 10%AF nf</td><td>9.9</td><td>1.2</td><td>23.7</td><td>1.2</td><td>10.6</td><td>27.2</td><td>1.7</td><td>30.7</td><td>nd</td><td>2.6</td><td>nd</td><td>nd</td><td>1.1</td></tr><tr><td>+1%T + 10%AF f</td><td>7.6</td><td>0.8</td><td>17.7</td><td>1.0</td><td>8.6</td><td>22.9</td><td>1.91</td><td>17.7</td><td>3.6</td><td>1.2</td><td>0.8</td><td>18.1</td><td>5.9</td></tr><tr><td>+2%T + 20%AF nf</td><td>14.4</td><td>1.8</td><td>25.8</td><td>1.7</td><td>12.8</td><td>31.1</td><td>1.8</td><td>22.2</td><td>nd</td><td>2.0</td><td>Nd</td><td>nd</td><td>1.1</td></tr><tr><td>+2%T + 20%AF f</td><td>11.4</td><td>1.1</td><td>19.6</td><td>1.3</td><td>11.6</td><td>24.2</td><td>2.0</td><td>13.8</td><td>4.0</td><td>1.0</td><td>0.9</td><td>15.6</td><td>5.9</td></tr><tr><td>+3%T + 30%AF nf</td><td>23.4</td><td>1.3</td><td>26.1</td><td>1.6</td><td>16.0</td><td>33.3</td><td>1.8</td><td>17.1</td><td>nd</td><td>1.6</td><td>nd</td><td>nd</td><td>0.9</td></tr><tr><td>+3%T + 30%AF f</td><td>17.0</td><td>1.0</td><td>19.8</td><td>1.3</td><td>13.3</td><td>25.9</td><td>2.0</td><td>13.6</td><td>3.7</td><td>1.0</td><td>0.8</td><td>12.6</td><td>5.1</td></tr></tbody></table>
### 3.5. Arachidonic Acid Yield

Comparing the total fatty acid profile of the fermented bioproducts we made the following observations (Table 4): (i) the percentage of ARA of total fatty acid content in bioproducts derived from the CM single- and two-component substrates was higher than for WB-based bioproducts; (ii) the total fatty acid profile of bioproducts derived from AF containing substrates was strongly affected by the fatty acid composition of the added AF—higher content of palmitic and stearic acid provided lower content of linoleic acid; (iii) the addition of the AF by-product to the SSF substrates resulted in the decrease of ARA content in the fermented BPs, where the degree of decrease was higher in CM-based bioproducts than in WB-based bioproducts. The levels of ARA in total fatty acids of bioproducts derived from CM-based substrates dropped from 26.5% (average value of single- and two-component substrates) to 8.1%, depending on the AF addition. On the other hand, the percentage of ARA of total fatty acids derived from WB-based bioproducts varied from 21.4% (average of single- and two-component substrates) to 12.6%, depending on the AF addition (Table 3). Comparing of bioproducts derived from single- and two-components substrates, it is clear that CM-based substrates were much more suitable for ARA biosynthesis than WB-based substrates (Figure 2). The average yield of ARA achieved using CM-based substrates was 36.2 mg ARA/g of BP (136.9 mg of ARA/g of fungal biomass), that is almost five times higher than the content of ARA in WB-based bioproducts, where the average yield from single- and two-component bioproducts was 7.5 mg ARA/g of bioproducts (42.2 mg ARA/g of fungal biomass). The addition of AF affected the ARA yield significantly (Figure 2). It caused a gradual cessation of ARA yield in CM-based bioproduct from 32.1 mg of ARA/g of bioproduct (addition of 5% (w/w) of AF) down to 21.2 mg ARA/g BP (addition of 30% (w/w) of AF). However, the content of ARA in bioproducts from the three-component CM-based substrates was at least double comparing to single- and two-components substrates. On the other hand, the addition of AF increased the yield of ARA in WB-based bioproducts up to 21.4 mg ARA/g of bioproduct (234.4 mg ARA/g of fungal biomass) in comparison to the fermented bioproducts from single-component substrates (7.1 mg ARA/g of bioproduct; 36.2 mg ARA/g of fungal biomass (Figure 2)).

<figure><img src="image_87.png" /><figcaption>**Figure 2.** Content of arachidonic fatty acid (ARA) in fermented bioproducts (BP) and in fungal biomass (FBM). T—Tween 40, AF—animal fat by-product.</figcaption></figure>

### 3.6. Biochemical Profile of the Fermented Bioproducts Obtained by FTIR-ATR Spectroscopy

FTIR-ATR spectroscopy allows rapid and non-destructive analysis of the total biochemical composition of different biological materials; in our case—bioproducts obtained by SSF of *M. alpina*.
Principal component analysis (PCA) of obtained FTIR-ATR spectra was used to study variations in the biochemical profile of the fermented bioproducts. The PCA scatter plot of the lipid-related spectral regions (3050–2800 cm⁻¹ and 1800–1700 cm⁻¹) shows significant chemical differences in the lipid profile for the individual fermented bioproducts obtained in SSF of substrates with various addition of AF (Figure 3). It was observed that the FTIR lipid profile of bioproducts obtained from the WB-based substrates was quite similar to the lipid profile of bioproducts derived from the substrates with 5% and 10% (w/w) of AF, while the lipid profile of bioproducts from the substrates with 20% and 30% (w/w) of AF showed a very different lipid profile (Figure 3). On the other hand, in case of CM-based bioproduct, PCA score plot (Figure 3) shows large differences between all tested conditions. The protein and polysaccharide profile of the fermented bioproduct from the SSF of AF supplemented substrates was similar, while it was different from the control substrates (Figure 3). The complete biochemical profile of bioproducts from the SSF of the control CM substrates was more similar to the CM substrates supplemented with 5% and 10% (w/w) of AF than in the case of WB substrates (Figure 3).

<figure><img src="image_88.png" /><figcaption>**Figure 3.** PCA scatter plots of FTIR-ATR spectra of fermented bioproducts obtained from the SSF of cornmeal-based and wheat bran-based substrates supplemented with animal fat at concentrations (blue) 0%, (red) 5%, (green) 10%, (orange) 20% and (yellow) 30%. (A)—lipid region of cornmeal-based fermented substrates, (B)—protein region of cornmeal-based fermented substrates, (C)—polysaccharide region of cornmeal-based fermented substrates, (D)—lipid region of wheat bran-based fermented substrates, (E)—protein region of wheat bran-based fermented substrates, (F)—polysaccharide region of wheat bran-based fermented substrates.</figcaption></figure>
4. Discussion

Oleaginous fungus *Mortierella alpina* is a well-known and thoroughly described species with a high capacity for lipid accumulation and a good ability to produce industrially relevant essential PUFAs [19–21]. In order to verify growth ability and utilization of AF by-products by *M. alpina* in SSF, two types of cereal matrixes (wheat bran and cornmeal) were used in this study. Cereal-based matrixes are easily accessible and well-known as a good matrix for SSF substrates. Due to their chemical composition, such as presence of easily accessible carbon, organic nitrogen and other macro- and micro-nutrients, cereal-based matrixes provide a useful source for good fungal proliferation, hyphae penetration and stable lipid accumulation in fungal cells [10,15]. In order to improve the conversion of cereal-based carbon into PUFAs enriched bioproducts, supplementation with exogenous oils containing precursors of PUFAs is advised. Thus, the addition of vegetable oils, such as sunflower, rapeseed, corn, soybean or linseed oil containing individual fatty acid precursors of PUFAs, led to a rapid increase of PUFAs yield in the final fermented bioproducts obtained by fungal SSF [10]. It was also reported that *M. alpina* is able to incorporate and transform exogenous fatty acids [22]. In this study, an addition of animal fat by-product as a source of PUFA precursors (such as palmitic acid, stearic acid or oleic acid) to the cereal-based matrix was evaluated for the production of ARA-enriched fermented bioproducts in an SSF by *M. alpina* CCF2861.

Due to the fact that cereal-based matrixes have a hydrophilic nature while animal fat is hydrophobic, there is a need to perform pretreatment of the fat materials before mixing it with cereal matrixes for obtaining homogenous substrates for SSF. The pretreatment of the AF by-product was performed by preparing homogenous fat-water-Tween®® 40 emulsions, where emulsifier Tween®® 40 forms a film around the dispersed lipid droplets and thereby reduces interfacial tension [29].

While the total carbon content in both substrates is very similar (37.4% for CM and 37.9% for WB), the difference is in the profile of the dominant carbon source. The main carbon source in cornmeal is starch accounting for up to 85% (w/w) of the total carbon, whereas wheat bran contains only up to 24.5% of starch and up to 50% of cellulose and hemicellulose [38,39]. *M. alpina* is known to effectively use starch as a carbon source, however the cellulose and hemicellulose polymers are not suitable source of carbon for this strain [40]. This might be a reason for better fungal growth of *M. alpina* on CM in contrast to WB. Analysis of protein content using the automatic Kjeldahl method for nitrogen content showed that WB contained higher amount of proteins (17.4%) than CM (6.5%), indicating lower C/N ratio in WB than in CM. However, the amount of proteins was sufficient for fungal growth and proliferation in SSF process for both substrates.

Nevertheless, as mentioned above, fungal growth in the presence of AF was undoubtedly affected mostly by the carbon structure (CM—starch, WB—cellulose and hemicellulose polymers), significant changes in pH values were also noticed. Subsequent TLC analyses of lipid classes of extracted total lipid clearly indicated that AF contains relatively higher levels of free fatty acids (Table 1), which was probably the reason for lower pH of the fermented bioproducts after cultivation with AF addition.

Fatty acid analysis of all fermented bioproducts obtained in this study confirmed the previously described good ability of *M. alpina* to produce ARA (Figure 2, Table 3) [4,20]. High ARA yield in the fermented bioproducts obtained after SSF of AF supplemented substrates proves a high potential of applying *M. alpina* for the biotransformation of AF by-products into high-value ARA-enriched fermented bioproducts.

FTIR-ATR spectroscopy performed on all obtained fermented samples has proved possible high potential of this method for analysis of different types of materials. This method requires low sample amounts and does not involve extraction of any chemical components [26]. The infrared (IR) spectrum contains information about chemical bonds characteristic to all major biomolecules of the measured sample and different spectral regions representing information about lipids, proteins, phosphate containing molecules and polysaccharides. When analyzing cellular lipid profile based on FTIR spectra, the most important lipid associated peaks, used for the analysis, are: (i) peaks related to -CH₃ and -CH₂ stretching at 2947 cm⁻¹, 2925 cm⁻¹, 2855 cm⁻¹, 1465 cm⁻¹ and 1377 cm⁻¹, indicating
mainly the chain length of the carbon skeleton in lipid molecules; (ii) the peak related to the ester bond stretching at 1745 cm⁻¹, indicating the total lipid content in the cell; (iii) the peak related to the carboxyl bond vibrations in free fatty acids at 1710 cm⁻¹, and (iv) the peak related to =C-H stretching at 3010 cm⁻¹, indicating the unsaturation level of cellular lipids. Proteins are observed in the spectral region 1700–1500 cm⁻¹ with the main peaks for amide I (1650 cm⁻¹) and amide II (1540 cm⁻¹) bonds and polysaccharides are observed in the region 1200–900 cm⁻¹. PCA of FTIR-ATR spectra proved distinct variations between individual samples in accordance with different composition of analyzed fermented samples. PCA score plots show large difference between all tested conditions, which could be caused by different composition of cereal substrate itself, as mentioned above. Higher similarity of complete biochemical profile in CM control samples with the CM substrates supplemented with 5% and 10% (w/w) of AF than in the case of WB substrates can be explained by the higher homogeneity of fungal growth and various cereal-based substrate utilization for the WB-based and CM-based substrates.

This is the first study on effective utilization of AF by-product into functional high-valued fermented bioproducts by SSF and obtained results represent broad fundamentals for subsequent research. Optimization of such fermentation process and extension for application of different fungal strains can lead to increased yields of desired metabolites or formatting a various high-value fungal specific product. Moreover, successful application of ATR-FTIR spectroscopy for rapid analysis of SSF bioproducts has been described, which can possibly extend industrial potential of the whole solid-state fermentation technology.

**Author Contributions:** Conceptualization, T.K.; methodology, O.S., T.K.; software, B.Z.; validation, T.K.; formal analysis, O.S., T.K., V.S.; investigation, O.S., T.K.; resources, T.K.; data curation, O.S., B.Z.; writing—original draft preparation, O.S.; writing—review and editing, T.K., V.S., M.Č.; visualization, O.S., T.K., A.K.; supervision, V.S., A.K., M.Č.; project administration, A.K., M.Č.; funding acquisition, V.S., A.K., M.Č. All authors have read and agreed to the published version of the manuscript.

**Funding:** This research was funded by Ministry of Education, Science, Research and Sports, Slovakia, grant VEGA 1/0323/19 and Council of Norway-FMETEKN grant, project number 257622; BIONÆR grant, project number 305215; HAVBRUK2 grant, project number 302543/E40; MATFONDAVTALE grant, project number 301834/E50 and DAAD grant, project number 309220. The funder had no role in the study design, data collection, and interpretation, or in the decision to submit the work for publication.

**Acknowledgments:** Authors would like to thank to Marije Oostindjer, Norilia AS (Oslo, Norway) for providing the animal fat by-products for experiments. Also, we would like to thank to Marek Jašurek and Pavel Lapeš for help with SSF experiments and to Margaréta Guttenová and Simona Dzurenová for assistance with ATR-FTIR analysis of SSF bioproducts.

**Conflicts of Interest:** The authors declare no conflict of interest.

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Article

# Aqueous Two-Phase System Extraction of Polyketide-Based Fungal Pigments Using Ammonium- or Imidazolium-Based Ionic Liquids for Detection Purpose: A Case Study

Juliana Lebeau<fnref n="1" />, Thomas Petit<fnref n="1" />,<fnref n="2" /> ID, Mireille Fouillaud<fnref n="1" /> ID, Laurent Dufossé<fnref n="1" /> ID
and Yanis Caro<fnref n="1" />,<fnref n="2" />,* ID

Received: 14 October 2020; Accepted: 14 December 2020; Published: 18 December 2020

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**Abstract:** Demand for microbial colorants is now becoming a competitive research topic for food, cosmetics and pharmaceutics industries. In most applications, the pigments of interest such as polyketide-based red pigments from fungal submerged cultures are extracted by conventional liquid-liquid extraction methods requiring large volumes of various organic solvents and time. To address this question from a different angle, we proposed, here, to investigate the use of three different aqueous two-phase extraction systems using either ammonium- or imidazolium-based ionic liquids. We applied these to four fermentation broths of *Talaromyces albobiverticillius* (deep red pigment producer), *Emericella purpurea* (red pigment producer), *Paecilomyces marquandii* (yellow pigment producer) and *Trichoderma harzianum* (yellow-brown pigment producer) to investigate their selective extraction abilities towards the detection of polyketide-based pigments. Our findings led us to conclude that (i) these alternative extraction systems using ionic liquids as greener extractant means worked well for this extraction of colored molecules from the fermentation broths of the filamentous fungi investigated here; (ii) tetrabutylammonium bromide, [N4444]Br-, showed the best pigment extraction ability, with a higher putative affinity for azaphilone red pigments; (iii) the back extraction and recovery of the fungal pigments from ionic liquid phases remained the limiting point of the method under our selected conditions for potential industrial applications. Nevertheless, these alternative extraction procedures appeared to be promising ways for the detection of polyketide-based colorants in the submerged cultures of filamentous fungi.

**Keywords:** fungal pigment; natural colorant; extraction ability; marine fungi; *Talaromyces albobiverticillius*; aqueous two-phases system extraction; ionic liquids

## 1. Introduction

Improving sustainability and reducing the amounts of waste produced and discarded by chemical industries is a global challenge. The production of dyes is one of the industries which is tackling these issues, aiming to minimize these impacts by using viable and beneficial alternatives to both human health and the environment. The demand for natural colorants has been growing over the past two decades and is now becoming a competitive research topic for food, cosmetics, pharmaceutics and

<footnote n="1">¹ Laboratoire de Chimie et de Biotechnologie des Produits Naturels (CHEMBIOPRO), Université de La Réunion, F-97490 Sainte-Clotilde, France; juliana.lebeau@gmail.com (J.L.); thomas.petit@univ-reunion.fr (T.P.); mireille.fouillaud@univ-reunion.fr (M.F.); laurent.dufosse@univ-reunion.fr (L.D.)</footnote>

<footnote n="2">² Département Hygiène Sécurité Environnement (HSE), Université de La Réunion—IUT La Réunion, F-97410 Saint-Pierre, France</footnote>

<footnote>* Correspondence: yanis.caro@univ-reunion.fr</footnote>
textiles industries [1]. The dyeing industry is currently suffering from the rising cost of raw materials due to the increasing demand for eco-friendly pigments to replace synthetic dyes (such as azo dyes). This is particularly true for the red-dye industries, which have no, or very few, natural red pigment alternatives with an adequate robustness meeting both the organoleptic expectations of consumers and the requirements of industrial processes. Unlike synthetic dyes, natural colorants are environmentally friendly as they do not require heavy and time- and energy-consuming benzene-based chemical synthesis, nor are they generating pollution and toxicity to human health wastewaters. Moreover, natural dyes often come as liquid solutions, while synthetic dyes tend to be dry powders, consequently requiring large amounts of water for dying purposes [2].

Recent literature has underlined the need to investigate new bio-pigments derived from bio-sources and notably pigments from microorganisms [3]. Indeed, the positive biological properties of some natural dyes, such as carotenoids and polyketide-based pigments of microbial origin, have made it possible to widen the potential area of specific applications of these natural colorants (such as for functional finishing of textiles, in functional foods, pharmaceuticals, etc.) [3]. Among unconventional sources, ascomycetous fungi of either marine or terrestrial origin are known to produce an extraordinary range of biomolecules such as bioactive polyketide pigments, which often tend to be more stable and water-soluble than plant pigments [4]. Thus, fungal colored polyketides are the most promising material in this respect. The production of natural colorants with a polyketide structure from filamentous fungi has already been reported, mostly using *Monascus*, *Talaromyces*, *Emericella*, *Paecilomyces* and *Trichoderma* species [3,4]. These aforementioned fungal species produce polyketide pigments with various hues, from yellow to reddish-purple, in the mycelium or as diffusing colorants in the fermentation broth (submerged culture). These fungal metabolites are structurally related (i.e., are biosynthesized by polyketide synthases, PKS) and are either azaphilone, naphthoquinone or hydroxyanthraquinone polyketide-based pigments [4].

However, to fully meet the demand of natural red colorants for the industries, there is an urgent need to develop and design green and sustainable detection, extraction and isolation techniques with higher recovery and greater selectivity in a faster and more benign manner than conventional methods. Indeed, the natural pigments available from plant, insect or microbial origins are currently essentially extracted using time-consuming, conventional liquid-liquid extraction methods that also require large volumes of various organic solvents. These conventional techniques usually involve prolonged contact with excess of organic solvents or a mixture thereof under high temperatures and/or pressures [5]. Moreover, these techniques sometimes require toxic solvents such as chlorinated or halogenated solvents, such as dichloromethane or toluene [6], and may impact on the final application (food, cosmetics, textiles, etc.) of the extracted colorants in regards to health and safety concerns. Furthermore, they often result in a relatively low extraction yield, poor selectivity and the need for further energy-intensive purification and wastewater recycling steps. Thus, any large-scale application raises environmental concerns, as well as uncertainties about their economic viability. Both scale-up and cost-effectiveness of the production of natural pigments at the industrial level, with a particular emphasis on red fungal pigments for food, cosmetic, pharmaceutical and fabric applications, still remain limited and require substantial optimization [4]. Ionic liquids, which are considered in this study, are new, unconventional solvents that have been receiving growing attention since the early 2000s because of their potential as green solvents in addition to their properties that are tunable to the target products and recyclability [7–13].

The aim of this study is the investigation of the potential of ionic liquids (ILs) as a more sustainable extraction technique for detection purpose of polyketide pigments from fermentation broths of filamentous fungi. Thus, we investigated the use of three different ammonium- or imidazolium-based ionic liquids, namely tetrabutylammonium bromide ([N4444]Br⁻), 1-butyl-3-methylimidazolium bromide ([C4Mim]Br⁻) and 1-butyl-3-methyl-imidazolium chloride ([C4Mim]Cl⁻), as new types of solvents in aqueous two-phase system (ATPS) extraction. An IL-ATPS is formed by mixing a soluble IL (chaotropic agent) and a salt with a kosmotropic profile (Na₂CO₃ and Na₂CH₃CO₂). These systems
allow the separation of two water-soluble compounds into two immiscible aqueous-rich phases. These specific ILs and salts were selected based on their water-miscibility profiles as well as their safety. It is worth mentioning that IL-based systems exhibit varying degrees of eco-toxicities driven both by the aromatic/alkyl chain of the IL itself as well as by the type of anionic salt it has been combined with [7,14,15]. As an example, tetrafluoroborate and hexafluorophosphate anions have been the commonly used salts with ILs due to their ability to enhance phase stability among other physicochemical advantages. However, fluorinated-based salts have the deleterious tendency to be hydrolyzed into toxic hydrofluoric acid [15]. Similarly, even the most hydrophobic ILs can be partially dispersed in water and become a potential polluting agent if not closely monitored. Only a few studies have addressed the environmental impact or potential risks associated with ILs' design, use and recycling. Thus, a rigorous selection of ILs and salt families and structures must be carried out beforehand based on the type of target biomolecules to be extracted, in order to make the most sustainable choice [14,16–18]. Imidazolium-based ILs such as [C4Mim]Br⁻ and [C4Mim]Cl⁻ are the most frequently used IL cores and are, therefore, well documented [19,20], hence we chose to study these in association with harmless salts. They were proven to have higher hydrophilicity than tetra-alkyl ammonium-based ILs ([N4444]Br⁻) because of their aromatic characteristics allowing a stronger interaction with water due to the delocalization of the positive charge over the aromatic ring [21]. However, the combination with anions and kosmotropic salt also has a significant impact on the final ability to form ATPS [22–24], as well as on the polarity of the overall system, and thus potentially influences the final partition of fungal pigments. Moreover, the extraction capabilities of the IL-based system will be challenged by the fact that the target fungal pigments are contained in fermentation broths. These are complex biological matrices containing a myriad of other components such as proteins, lipids and other excretable secondary metabolites, which can most likely impact the chemical structure and availability of the target compounds. Therefore, both hydrophilic and hydrophobic IL cations combined with ATPS-forming anions and organic and inorganic salts were investigated here.

Within this scenario, we investigated this alternative IL-ATPS extraction technique using the three aforementioned ammonium- or imidazolium-based ionic liquids to selectively extract the pigmented polyketides from submerged cultures of four filamentous fungi producing a mixture of pigments, namely *Talaromyces albobiverticillius* (producing deep red pigments), *Emericella purpurea* (producing red pigments), *Paecilomyces marquandii* (producing yellow pigments) and *Trichoderma harzianum* (producing yellow to brown pigments). These four strains were selected because they are well known to produce biomolecules exhibiting a rich chemical and structural diversity with different colors, including red ones [25,26]. *Talaromyces* represents a good example of the diversity of fungal pigments. In our previous study, we highlighted the potential of the marine-derived fungus *T. albobiverticillius* 30548 to produce, in a submerged culture, azaphilone polyketide-based red colorants, such as N-threonine-monascorubramine, N-glutaryl-rubropunctamine and PP-O [26]. In other previous studies, some azaphilone (hexaketide) compounds with biological activities have also been isolated from the fermentation broth of *T. harzianum*, such as the yellow azaphilone pigment T22 [27] and the compounds harziphilone and fleephilone [28], with some other extrolites with different chemical structures (cyclonerodiol, trichosetin, harzianic acid, harzianopyridone and MR566A) [29]. Some species of *Trichoderma* are natural producers of anthraquinone (octaketide) pigments, whose biological activities are also well known. For example, several anthraquinone pigments have been reported earlier from a fermentation broth of *T. harzianum*—crysophanol (red), pachybasin (orange), emodin (orange), ω-hydroxypachybasin, 1,5-dihydroxy-3-hydroxymethyl-9,10-anthraquinone, 1,7-dihydroxy-3-hydroxymethyl-9,10-anthraquinone, 1,8-dihydroxy-3-methylanthraquinone and 1-hydroxy-3-methylanthraquinone [30,31]. Catenarin (red) is an anthraquinone derivative that has been isolated from a variety of fungi, including species of *Emericella* [4]. *Emericella* sp. is also known for its ability to produce other bioactive secondary metabolites with different chemical structures, including polyketide pigments. In particular, various extrolites such as epurpurin A–C,
variecolins, versicolorins, albistrins, emerin, emindol PA, norsolorinic acid and shamixanthones have been isolated from *Emericella purpurea* [32]. Next, sorbicillinoids (e.g., sorbicillin, sorbicillinol, oxosorbicillinol and bisvertinol) are an important family of hexaketide compounds produced by terrestrial and marine fungi, including species of *Trichoderma* (e.g., *T. harzianum*) and *Paecilomyces* (e.g., *P. marquandii*) [33].

In regards to the diversity of the expected compounds, the selectivity of the IL-ATPS extraction for specific types of fungal pigments (e.g., red ones) was investigated. Thus, the main goal of our study here is the selective extraction of the fungal polyketide-based pigments with a red hue from the fermentation broths. Additionally, we looked into the reverse extraction of the colored molecules from the IL-ATPS by working on the ionizable profile of the polyketide molecules. Ultimately, this gave us tools to estimate the overall rate of pigment recovery from the fermentation broth and identify points of future improvements. It is worth mentioning that most of the studies conducted so far using IL-based extraction methods on biomaterials (fungal-, plant- or insect-originated) explored only either the extraction power of the IL on pure solution of compounds of interest or the stability and/or potential use of the said compounds in the IL phase [34]. Few and infrequent studies have been conducted on testing the recovery of the molecule of interest from the IL-based extractant as well as its reuse [34]. Nevertheless, it may be noted that the accurate composition of all the secondary metabolites produced by these aforementioned fungal strains has not been investigated here, because that is not the issue.

## 2. Materials and Methods

### 2.1. Selection of the Ionic Liquids and Formation of the Kosmotropic Salt Buffer

The compositions and structures of the three ionic liquids, ([N4444]Br⁻), ([C4Mim]Br⁻ and ([C4Mim]Cl⁻), used in this study are presented in Table 1. The reagents were purchased from Sigma Aldrich (Saint Louis, MO, USA). Alizarin (anthraquinone) purchased from Sigma Aldrich was used as control for IL-ATPS extraction purposes.

<table><caption>**Table 1.** Description of the selected ionic liquids (ILs) associated with the kosmotropic salt buffer, enabling a diphasic system.</caption><thead><tr><th>ILs</th><th>Salt</th><th>Structure</th></tr></thead><tbody><tr><td>[C4Mim]Br<sup>-</sup><br>1-Butyl-3-methylimidazolium bromide</td><td rowspan="2">Na<sub>2</sub>CO<sub>3</sub><br>Sodium carbonate<br>(Inorganic)</td><td>[Image of structure for [C4Mim]Br<sup>-</sup>]</td></tr><tr><td>[C4Mim]Cl<sup>-</sup><br>1-Butyl-3-methylimidazolium chloride</td><td>[Image of structure for [C4Mim]Cl<sup>-</sup>]</td></tr><tr><td>[N4444]Br<sup>-</sup><br>Tetrabutylammonium bromide</td><td>NaCH<sub>3</sub>CO<sub>2</sub><br>Sodium acetate<br>(Organic)</td><td>[Image of structure for [N4444]Br<sup>-</sup>]</td></tr></tbody></table>

### 2.2. Fungal Strains and Culture Conditions

The marine-derived fungal strain *T. albobiverticillius* (collection number 30548, GenBank accession number MK937814) was isolated from the back reef-flat of Réunion Island [35]. The three other strains used in this study were isolated from terrestrial environments, including *T. harzianum* (commercial biological control strain T22), *E. purpurea* (collection reference LCP 3323) and *P. marquandii* (collection reference LCP 2271) and were bought from the fungal culture collection of the Muséum national d'Histoire naturelle (MNHN, Paris, France). For submerged cultures, potato dextrose broth
(PDB; Sigma Aldrich, Saint Louis, MO, USA) was used routinely as culture medium. The pH of the culture medium was adjusted to 6.0 ± 0.2 using 0.1 M HCl prior to sterilization. Pre-culture and cultivation were carried out in 250 mL Erlenmeyer flasks containing 100 mL of sterilized culture medium. The flasks were incubated at 26 °C for 7 days and agitated at 150 rpm using a rotary agitator (Infors Multitron, 50 mm excentration, Infors HT, Switzerland) as described by Lebeau and coworkers (2017) [25]. After 7 days of fermentation, the contents of each flask were collected and centrifuged at 14,000× g for 10 min; the resulting supernatant was filtered through a Whatman filter paper (GF/C) at a reduced pressure using a Büchner funnel to obtain the culture filtrate. The filtered colored supernatants were then freeze-dried at -84 °C in an ultra-low-temperature freezer (Sanyo, Guangzhou, China) for at least 2 h. The samples were then quickly transferred to a LABCONCO FreeZone 2.5 lyophilizer (LABCONCO, Kansas City, MO, USA) and lyophilized for 24 h. During freezing, the condenser temperature and vacuum pressure were maintained at -47 °C and 200 mbar, respectively (Figure 1).

<figure><img src="image_93.png" /><figcaption>**Figure 1.** Brief description of the overall ideal IL aqueous two-phase system (IL-ATPS) workflow for direct detection and extraction of fungal pigments into ionic-rich phase, their back-extraction into salt-rich solution and re-injection of IL into the process.</figcaption></figure>

## 2.3. Binodal Curves Determination

Ternary phase diagrams for each of the IL-ATPS-forming systems were determined using the Cloud Point titration method at room temperature and pressure [36]. Aqueous solutions of salts at 40% *w*/*w* and aqueous solutions of ILs at 30% *w*/*w* were also prepared at room temperature and pressure and were used for the determination of the binodal curves for each IL-ATPS. It was carried out by the drop-wise addition of the aqueous solution of the concentrated salt to the IL aqueous solution (or pure polymer) until the detection of a cloudy and biphasic solution. This was followed by the drop-wise addition of pure water (diluent) until the formation of a clear and limpid solution corresponding to the monophasic regime. The composition of the two-phase system was determined using the ratio of the weight of one component added to the total weight of all added components. The above procedure was repeated until the least amount of turbidity was observed to obtain sufficient data to generate binodal curves [37,38]. At the level of the biphasic region, the upper phase was composed by the IL-rich aqueous phase while the lower phase was the salt-rich phase for all ILs and salts investigated in this study.

## 2.4. Fungal Pigments Partitioning Using Ionic Liquids in an Aqueous Two-Phase System Extraction

Ternary mixtures within the biphasic region were prepared for each IL-ATPS and consisted of either (1) 15 wt.% of [C4Mim]Br⁻, 15 wt.% Na₂CO₃ and 70 wt.% of the aqueous solution of pigmented fungal broth; (2) 15 wt.% of [C4Mim]Cl⁻, 20 wt.% Na₂CO₃ and 65 wt.% pigmented fungal broth; and (3) 15 wt.% of [N4444]Br⁻, 25 wt.% NaCH₃CO₂ and 60 wt.% pigmented fungal broth. The final volumes of the reaction mixtures were about 5 mL (final total weight of 5 g), with 0.75 g of the corresponding ionic liquid for 1 g of lyophilized fungal broth resuspended in corresponding weights
of water and salt (i.e., 2.5 g of water + 0.75 g of Na₂CO₃, 2.25 g of water + 1 g of Na₂CO₃ or 2 g of water + 1.25 g of NaCH₃CO₂, respectively). The ternary mixtures' compositions were chosen based on the phase diagrams of each IL investigated (Figure 2). The extraction step was performed as follows: all components (lyophilized pigment broth previously obtained by fermentations, salts, water and IL) were mixed together, vigorously stirred and allowed to equilibrate for 30 min before centrifugation at 2455× g (4000 rpm) for 5 min. The mixture was then allowed to decant overnight (16 h). The next day, each fraction of the mixtures was carefully separated, weighted out and its volume was measured. These values were used to calculate the coefficient of partition of pigments (Kₚigment), the phase ratio (R) and the selectivity of type of pigments ratio (Sₚigment), as well as the efficiency of extraction (Effp%), as described in the following sections.

<figure><img src="image_94.png" /><figcaption>**Figure 2.** Binodal curve for the determination of weight ratio of IL, salt buffer (Na₂CO₃ or Na₂CH₃CO₂) and water to produce biphasic systems. R1, R2 and R3 display the composition of the ATPS made with [C4Mim]Br⁻, [C4Mim]Cl⁻ and [N4444]Br⁻, respectively.</figcaption></figure>

The recovery of the pigments extracted from the IL phase was carried out by resuspending the previously collected IL phase in concentrated aqueous salt solution and adjusting the pH to 13. The mixture was stirred and left to stand for 30 min before being centrifuged at 2455× g (4000 rpm) for 5 min and allowed to decant overnight (16 h). Similar measurements of weight, volumes and absorbance were carried out and the efficiency of pigment recovery was calculated as described below.

## 2.5. Determination of the Coefficients of Partition of Pigments (Kₚigment)

After the extraction steps, samples of both IL-rich and salt-rich phases were diluted in distilled water and the pigment content of each phase was estimated by measuring the absorbance using a UV spectrophotometer (UV-1800, Shimadzu Corporation, Tokyo, Japan) at 490, 470 and 400 nm corresponding to red, orange and yellow hues, respectively. Relative repartitions (Kₚigment) of each type of pigment (red, orange and yellow) were calculated according to the following Equations (1)–(3):

$$K_{\text{red}} = (A_{490_{\text{IL}}} * \text{FD}_{\text{IL}})/(A_{490_{\text{salt}}} * \text{FD}_{\text{salt}}) \quad (1)$$

$$K_{\text{orange}} = (A_{470_{\text{IL}}} * \text{FD}_{\text{IL}})/(A_{470_{\text{salt}}} * \text{FD}_{\text{salt}}) \quad (2)$$

$$K_{\text{yellow}} = (A_{400_{\text{IL}}} * \text{FD}_{\text{IL}})/(A_{400_{\text{salt}}} * \text{FD}_{\text{salt}}) \quad (3)$$

where Kₚigment: coefficient of partition of the target shade of pigment (red, orange or yellow pigments); Aₗl: absorbance (mAU) of the IL phase sample at the wavelength considered (490, 470 or 400 nm); FDₗl: dilution factor of the IL phase sample; Aₗsalt: absorbance (mAU) of the salt phase at the wavelength considered (490, 470 or 400 nm); FDₗsalt: dilution factor of the salt phase.
## 2.6. Determination of the Selectivity of Type of Pigments Ratio (S<sub>pigment</sub>)

In order to evaluate the higher selectivity of ILs tested against the isolation of either red, yellow or orange pigments, the selectivity of type of pigments ratio (S<sub>pigment</sub>) was determined as follows:

$$S_{\text{red/yellow}} = K_{\text{red}}/K_{\text{yellow}} \qquad (4)$$

$$S_{\text{red/orange}} = K_{\text{red}}/K_{\text{orange}} \qquad (5)$$

where S<sub>red/yellow</sub>: selectivity ratio of red to yellow pigments extracted; K<sub>red</sub>: coefficient of partition of the red pigments extracted; K<sub>yellow</sub>: coefficient of partition of the yellow pigments extracted; S<sub>red/orange</sub>: selectivity ratio of red to orange pigments extracted; K<sub>orange</sub>: coefficient of partition of the orange pigments extracted.

## 2.7. Determination of the Extraction Efficiencies (Eff%)

The efficiency of the direct extraction was determined according to the following formula:

$$\text{EffD\%} = 100 \times [K_{\text{pigment}}/(1 + (1/R))] \qquad (6)$$

where EffD%: efficiency of the direct extraction; K<sub>pigment</sub>: coefficient of partition of the target pigments extracted; R: phase ratio between the IL- and salt-rich phase volumes.

The phase ratio (R) was calculated using the following equation:

$$R = V_{\text{IL}}/V_{\text{salt phase}} \qquad (7)$$

where R: phase ratio; V<sub>IL</sub>: volume (mL) of the IL phase sample; V<sub>salt</sub>: volume (mL) of the salt phase.

# 3. Results

## 3.1. Determination of Biphasic Systems Using Binodal Curves

In the search for alternative extraction systems that are less hazardous for the environment and for human health, the properties of two different families of ionic liquids, namely the imidazolium- and the tetra-alkyl ammonium-based core families, were investigated specifically for extracting mixtures of fungal pigments. Subsequently, three IL-ATPSs, including [C4Mim]Br<sup>-</sup> + Na<sub>2</sub>CO<sub>3</sub>, [C4Mim]Cl<sup>-</sup> + Na<sub>2</sub>CO<sub>3</sub> and [N4444]Br<sup>-</sup> + NaCH<sub>3</sub>CO<sub>2</sub>, were investigated to evaluate their ability to selectively extract and detect mixtures of fungal pigments with different colors from fermentation broths [25]. Fungal fermentation broths are complex biological matrices containing various complex compounds, from proteins to lipids and other excretable secondary metabolites, including fungal pigments. Such diversity represents a burden for the development of fast, efficient and cost-effective detection and extraction processes, for which IL-ATPSs are expected to offer realistic options [19,21]. A brief description of the overall extraction process is shown in Figure 1. In this study, IL-ATPSs were applied to a lyophilized pigmented fermentation broth where the fungal pigments we are searching for (e.g., polyketide-derived pigments such as azaphilone red compounds, etc.) should have hydrophilic profiles.

Imidazolium cores have a stronger interaction with water than tetra-butyl ammonium and, consequently, tend to attract more hydrophilic systems. The combination with different anions and salts has an impact on the final overall hydrophilic/hydrophobic profile of the ionic liquid-rich phase, and this can be assessed by the determination of the binodal curves of each system [37–39]. Thus, binodal curves were produced for each of the IL-ATPSs in order to determine which ionic liquid/salt ratio was needed to generate a biphasic system. The resulting curves were produced using the Cloud Point titration method [28,36] and are displayed in Figure 2. We designed three IL-ATPSs that have in common the same ionic liquid content (15%), with the underlying strategy to avoid a large
amount of ILs to be used due to both their cost and environmental impact. Although ionic liquids are considered as more ecological extraction solvents because of their higher extraction capacities for smaller volumes and recyclability, their final discard remains a grey area with regard to respect for the environment [14]. The corresponding amount of salt to be added for each specific ionic liquid system was established based on the binodal curves. Because clear and stable two-phase systems are desired, the less ATPS-forming a system is, the higher the amount of added salt (% *w/w*) was, in the limit of 25% *w/w* of the system. Both salts, sodium carbonate and sodium acetate, were adopted on the basis of their pH-buffering ability, as well as their biodegradability and lack of toxicity [40,41].

As shown in Figure 2, the phase-forming ability of the different ionic liquid systems can be ranked in the following order: [C4Mim]Br⁻ > [C4Mim]Cl⁻ > [N4444]Br⁻. From the binodal curves, surprisingly, the most ATPS-forming systems were the imidazolium-based ATPS. Due to its longer alkyl chains, [N4444]Br⁻ was expected to be the most prone to generate a two-phase systems. Among the systems investigated here, imidazolium-based ionic liquids were the least ATPS-forming options against tetra-alkyl ammonium-based ionic liquids, when taken under the same conditions (i.e., combination with the same anion and chaotropic/kosmotropic salts). Between the two classes of imidazolium-based ionic liquids chosen here, the only difference is the coupled anion (the salt being identical). [C4Mim]Br⁻ showed higher ATPS-forming capacity than [C4Mim]Cl⁻, and this is in line with previously reported results, where it was observed that bromide-based IL had greater aptitude to form ATPS than its chlorine anion-linked counterpart [37,42], when used under the same conditions (salts and ionic liquid core). This can be explained because bromide anions are more chaotropic than chlorine anions. The [N4444]Br⁻ system involves a different salt, namely an organic salt of sodium acetate. A previous study comparing the effect of the salt on the ability to generate a two-phase system reported sodium carbonate, an inorganic salt, as significantly inclined to form biphasic systems [37]. It is then reasonable to assume that the lower ability of [N4444]Br⁻ to form an ATPS, despite its longer alkyl chains, is mostly due to its combination with the organic salt of sodium acetate [43].

Those systems and their variation in profiles and behaviors have provided an unusual method for the selective extraction of pigments produced by submerged cultures of filamentous fungi from their fermentation broth. Indeed, we seek to selectively extract hydrophilic polyketide-derived pigments from ionic liquid-phase mixtures; therefore, we are investigating relatively hydrated ionic liquid phases that are still capable to form stable ATPSs.

## 3.2. *Effect of the Ionic Liquids Aqueous Two-Phase System Extraction in the Partitioning of Fungal Pigments*

In a previous published study, we identified the following four fungal strains: *T. albobiverticillius* of marine origin (deep red) and three strains of terrestrial origin, i.e., *E. purpurea* (red), *P. marquandii* (yellow) and *T. harzianum* (yellow-brown), based on their capacities to produce high amounts of polyketide-derived pigments in fermentation broth. One of the main limiting steps in the bioproduction of pigments is their inefficient and cost-effective extraction. We selected the aforementioned fungal strains to investigate the potential of IL-ATPS extraction for the selective detection and extraction of fungal pigments with different colors. Currently, there is a specific need for natural alternatives for red pigments for industrial use. We therefore focused our work on the potentialities of extracting fungal red pigments by IL-ATPS selected here and, more particularly, on the azaphilone-like red pigments from the fermentation broth of the marine-derived filamentous fungi *T. albobiverticillius* 30548, which was collected on the west coast of Réunion Island. Both tetra-butyl ammonium and imidazolium ionic liquids were chosen because they are considered more compatible (and less disruptive) with biomolecule structures and have proven large spectra of applications. Moreover, previous studies reported successful partitioning and recovery of hydrophobic pigments such as carotenoids using IL-ATPSs with similar polarity profiles as the target compounds [21,22,40,44,45]. Thus, the same strategy was applied here by investigating IL-ATPSs with greater hydrophilic profiles for the purpose of better azaphilone-like-pigment isolation. Based on these initial properties, it was assumed that the more hydrophilic the IL top phase, the higher the pigment partition coefficient and, consequently,
the better the extraction would be. The results of the first direct extraction are shown in Figure 3. Globally speaking and across all three ATPSs tested here, the results of extraction showed that pigments tend to preferentially migrate towards the top ionic liquid-rich phase. Unanimously, [C4Mim]Br⁻ with 15 wt.% Na₂CO₃ exhibited the smallest pigment partition in the four pigment-producing strains. Indeed, there was a significant loss of pigments remaining in the aqueous salt-rich phase, as shown in Figure 3. On the other hand, both systems, i.e., [C4Mim]Cl⁻ + 20 wt.% Na₂CO₃ and [N4444]Br⁻ + 25 wt.% NaCH₃COO⁻, showed significantly higher pigment migration in the IL top phase, confirming their greater potential as an extraction system for hydrophilic fungal polyketide-derived pigments of various colors (red, orange and yellow).

<figure><img src="image_95.png" /><figcaption>**Figure 3.** Visual aspect of the first step of extraction from the different IL-ATPSs tested and their extraction abilities on fungal fermentation broth and pure pigment control (alizarin). Lyophilized colored fermentation broths were mixed with the corresponding ionic liquid and salt. The mixtures were allowed to equilibrate for 30 min before centrifugation for 5 min at 4000 rpm and overnight decantation. Both ionic liquid-rich and salt-rich phases were collected and absorbances were measured. Top phase: ionic liquid-rich phase where most of the pigments migrated; Bottom phase: salt-rich phase.</figcaption></figure>

The coefficients of partition of distinct pigment shades ($K_{red}$, $K_{orange}$ and $K_{yellow}$) express the preference of each type of pigment to migrate in ionic liquid-rich phase. They were then calculated for each system and are displayed in Figure 4. In line with previous observations, [C4Mim]Br⁻ + 15 wt.% Na₂CO₃ resulted in the lowest K values for all pigment shades for the four strains (Figure 4) and it was concluded as a non-suitable system for our purpose. Although the partition coefficient values obtained from the two different remaining ATPSs on the same fermentation broths suggested that the design of the IL-based ATPSs has an impact on pigment migration, it is important to consider that the composition in pigments and their respective polarity profiles will have an impact on the efficiency of the overall extraction of the color shade. We therefore investigated two red-pigment-producing fungal strains, namely *T. albobiverticillius* (30548) and *E. purpurea* (3323), which were submitted to the same
extraction process. Interestingly, [N4444]Br⁻ + 25 wt% NaCH₃COO statistically showed significantly better pigment partition coefficients for red to orange pigments from *T. albobiverticillius* than those from *E. purpurea* and *T. harzianum*. On the other hand, the extraction efficiency was equally as good for both strains using [N4444]Br⁻- and [C4Mim]Cl⁻-based ATPSs (Figure 5). These observations led to the conclusion that quarterly ammonium-based IL-ATPS (i.e., N4444]Br⁻) has better extraction capacities towards azaphilone-like red pigments. Moreover, it was noticed that the extraction yields tended to be less repeatable across the replicate experiments performed on pigments from *E. purpurea* for both [N4444]Br⁻ and [C4Mim]Cl⁻ based ATPSs. This led to the conclusion that the composition of the pigment mixture highly influenced the efficiency of one system. Previous studies have concluded on the production of azaphilone-like red compounds mainly by *T. albobiverticillius*, suggesting mostly hydrophilic molecules. [26]. The binodal curve initially established here highlighted the [N4444]Br⁻-ATPS as the most hydrophilic system amongst our ATPSs, which could explain why and how this specific system appeared to be a more appropriate extraction agent for an apparent homogenous mixture of reddish hydrophilic pigments.

<figure><img src="image_96.png" /><figcaption>**Figure 4.** Partition coefficients of red (490 nm) (A), orange (470 nm) (B) and yellow (400 nm) (C) pigments for each strain in the four IL-ATPSs tested (*, ** and *** stand for statistical significance based on *p*-values < 0.05 performed for the same strain treated with different IL-ATPSs, *n* = 3).</figcaption></figure>
<figure><img src="image_97.png" /><figcaption>**Figure 5.** Direct extraction efficiency of red (490 nm) (A), orange (470 nm) (B) and yellow (400 nm) (C) pigments for each strain in the four IL-ATPSs tested (* indicate *p*-values < 0.05, *n* = 3).</figcaption></figure>

In parallel, the repartition coefficient for yellow pigments remained drastically smaller than repartition coefficient for red, K<sub>red</sub>, across all ATPSs as well as amongst all fungal strain productions. Even the bright-yellow-pigment-producing fungal strain, namely *P. marquandii*, did not generate partition coefficient values as high as red-pigment-producing fungal strains (i.e., *T. albobiverticillius* and *E. purpurea*) for red to orange pigments. Such limited results for yellow compounds could be explained by the fact that reddish and orangish pigment-producing fungal strains may simply not produce many actual yellow molecules. In the case of the strong yellow-producing strain of *P. marquandii*, although the pigments clearly migrated to the ionic liquid-rich phase, the raw data of absorbance
measured at 400 nm (yellow typical wavelength) were globally four times weaker in reddish strains, suggesting a smaller inherent yellow-pigment-production compared to red. Nevertheless, as shown in Figure 4C, no statistically significant differences were observed in yellow compound extractions from one IL-ATPS to another when used on broth containing a mixture of pigments produced by *T. albobiverticillius*, *E. purpurea* or *T. harzianum*. On the contrary, on the yellow-producing strain, namely *P. marquandii*, a statistically significantly better extraction capacity (i.e., higher K<sub>yellow</sub>) was observed with [C4Mim]Cl<sup>-</sup>.

Thus, from this first round of extraction, [N4444]Br<sup>-</sup> appeared as the more appropriate system amongst the ones investigated herein for consistent and efficient extractions of azaphilone-like red compounds from fungal resources. The yellow polyketide-based secondary metabolites produced by the investigated fungal strains were not the most preferred pigments extracted by either [N4444]Br<sup>-</sup> or [C4Mim]Cl<sup>-</sup>; however, [C4Mim]Cl<sup>-</sup> showed relatively good robustness and efficiency with respect to the extraction of these yellow compounds (such as yellow azaphilone pigment T22 from *T. harzianum* and yellow sorbicillinoids (hexaketide compounds) from *P. marquandii*). It is worth mentioning that it seems more suitable to use this system on a strain generating a strongly yellow broth, as more inconsistent extraction results were observed on the yellow-brown mixture produced by *T. harzianum*. Thus, further optimizations, in particular on salt combinations and/or ratio thereof, could lead to better detection and extraction capacities.

### 3.3. Selectivity of the IL-ATPS for Red Pigments in Various Fungal Broths

As mentioned earlier, the selective isolation of red pigments from other fungal pigments and cellular components is the main goal of this study, due to the higher potential market of red colorants for industrial application [3,4,25]. To further determine the ability of each IL-ATPS to detect and isolate red colorants, two selectivity parameters were calculated as the ratio of the partition coefficients of red pigments to other fungal pigments and are displayed in Figure 6.

<figure><img src="image_98.png" /><figcaption>**Figure 6.** Extraction selectivity of red pigments over yellow pigments, S<sub>red/yellow</sub> (A) and over orange S<sub>red/orange</sub> (B) pigments for each strain in the four IL-ATPSs tested (* indicates p-values < 0.05, n = 3).</figcaption></figure>
Overall, in red-producing strains, the selective extraction of red over yellow co-produced colorants was strongly favored, as suggested by the high S<sub>red/yellow</sub> values (i.e., ratio S<sub>red/yellow</sub> >>1) reported across the IL-ATPSs considered. In particular, we reported a statistically significantly better selectivity of red pigments' extraction from *T. albobiverticillius* when using [N4444]Br<sup>-</sup> compared to [C4Mim]Cl<sup>-</sup>, while no significant difference was observed from *E. purpurea*. This suggested that the migration and selectivity of mixtures of pigments are greatly impacted by the features of the ILs used. As far as we can see and without knowing the detailed composition of pigments in the different mixtures, we confirmed that the [N4444]Br<sup>-</sup>-based ATPS was the most suitable extraction system for the selective isolation of red colorants across different hue-producing strains. On the other hand, little or no selection occurred between red and orange pigments as shown in Figure 6B and in all IL-ATPSs and studied strains. Indeed, we reported a selective extraction ratio for red/orange pigments close to 1 for all selected strains (Figure 6B). We also noticed small distinction between red and orange or yellow colorants in all the IL-ATPSs used on *T. harzianum* (Figure 6A,B), suggesting a strong likeness of concomitant extraction of red and orange pigmented molecules from the fermentation broth showing a mixture of pigments of different shades. These observations led to the conclusion that our [N4444]Br<sup>-</sup> system showed its highest potential under the conditions investigated here when applied on single-tone broth.

### 3.4. Reverse Extraction of the Fungal Pigments from the IL-ATPSs

The main objective of this study was to explore the potential of IL-ATPSs for the selective detection and extraction of red fungal pigments from a mixture of colored molecules. We succeeded in showing promising detection and extraction efficiencies of the [N4444]Br⁻ system for azaphilone-based compounds. This satisfied our initial scientific question. To further explore the potential applications of this technique, we investigated the back-extraction efficiency of the pigments from the IL system using a previously reported method. This was carried out with the purpose of rapidly estimating how straightforward and repeatable back extractions from IL-based systems across different types of biological matrixes are. Although several publications have reported the benefits of using IL-based extraction systems for the isolation of bioactive compounds [46], very few have addressed the corresponding challenges to develop efficient extraction of the said bio-compounds from IL-based solvents (referred to as back-extraction) [23,47]. Tan and coworkers [43] succeeded in back-extracting the hydrophilic anthraquinones of aloe-emodin from their IL-ATPS with [C4Mim]BF4 by adding an alkaline salt solution onto the IL-rich phase containing the pigment. The same approach was applied here using an alkaline (pH 13) solution of the respective salts initially used for each system, which were added to the separated IL-rich phase, shaken and left for decantation overnight. The visualization of the back-extraction steps before and after the alkalinization of the salt solution is shown in Figure S1. The visual aspects of the extracts (Figure S1) indicate that the reverse extractions remained extremely limited. The increase in pH up to 13 using drop-wise addition of NaOH did not allow a better recovery of any of the pigments tested under our experimental conditions. After the addition of the salt solution and before the alkalinization of the system, the phase separation between the IL- and salt-rich phases became blurry, especially for the [C4Mim]Cl⁻ system (Figure S1). Interestingly, in the [C4Mim]Cl⁻ and [C4Mim]Br⁻ ATPSs, before the pH increase, a slight repartition of the pigments in the salt phase for the red-producing strains (*T. albobiverticillius* and *E. purpurea*) and the brown-producing strain (*T. harzianum*) was observed. The increase in pH appeared to have counteracted the recovery of the pigments in the salt-rich phase, whilst further improving the partition in the IL-rich phase, as can be seen in particular with the clarification of the salt-rich phase in the case of the [C4Mim]Cl⁻ across the four fermentation broths tested (Figure S1).

The overall efficiencies and abilities of recovery of red pigments from the IL-ATPS after application of an alkaline salt solution are displayed in Figure S2. We concluded that the recovery of the pigments from any of the fermentation broth tested was very poor under these conditions. The highest overall recovery was obtained from the fermentation broth from *E. purpurea*, with a final recovery of about 20% (Figure S2B). We were also forced to conclude that we could not extract back, from the IL phase,
the dark red pigments excreted by either *T. albobiverticillius* or *E. purpurea* because the pigments did not migrate to the alkaline salt-rich phase. Curiously, it seemed that increasing the pH reduced the migration of the pigments, which was in contradiction with the results obtained by Ventura et al. [40] on hydrophilic anthraquinone of aloe-emodin. However, this can be put into perspective by the fact that, firstly, we do not know the exact hydrophilic/hydrophobic equilibrium of all the fungal pigments produced in the fermentation broths, and secondly, we directly applied these IL-ATPSs onto complex fermentation matrices. Then, plenty of other metabolites were likely to interfere in the selective extraction of pigmented compounds. Globally speaking and despite the low overall recovery values, we could notice a statistically significant difference in the [N4444]Br⁻-based IL-ATPS on the extraction, back-extraction and overall recovery of red pigments. In conclusion, we were able to recover more red pigments from the fermentation broth of the second most red-pigment-producing strain, namely *E. purpurea*, by using the [N4444]Br⁻-ATPS.

To date, ionic liquids are considered to be a sustainable means of extraction, mainly because of their very high extraction power, perfectly illustrated here, revealing a promising potential as cost-effective, profitable and less hazardous solvents. However, it is important to recall that ionic liquids require heavy organic chemical synthesis, which can be counterintuitive against their reputation of being new-age green solvents. However, few arguments can be given here in favor of ionic liquids' green profile against conventional extraction solvents. First, the already well-known higher extraction power and recyclability of ionic liquids are their dire assets, as previously said. Second, a recent paper estimated and compared the environmental sustainability between a similar IL-based extraction system applied to carotenoid recovery and a conventional solvent extraction method (using acetone and ethanol) by calculating environmental factor (E-factor) and carbon footprint parameters [45]. The E-factor and carbon footprint calculations included all greenhouse gas (GHG) emissions from the production of all chemicals and the water and electricity consumed during the process. It was concluded that IL-based extraction offered a significantly reduced environmental impact compared to conventional solvents. Perspective must be taken though, as efficient recycling of both IL materials and water is paramount for ionic liquids to be properly referred to as green solvents.

The poor overall recovery obtained here highlights essential points of improvements to be made while perfectly illustrating how much case-by-case optimization is needed to use ionic liquids for detection and extraction purposes. It is worth mentioning that there are various options to explore in order to optimize back-extractions and enable multiple re-use of ionic liquids, such as using a different salt or polymer solutions as antisolvents, changing temperatures and applying ultrasounds or microwaves and a combination thereof [48]. Depending on the type and sensitivity of the target compounds trapped in IL phase, distillation or even adsorption can also be used. However, these options need to be carefully considered based on the additional energy, water, time and specific equipment expenditures they would require. However, this point was not in the scope of this paper.

4. Conclusions

Our findings suggested that the combination of an ATPS extraction system using ionic liquids as greener extraction solvents is a possible tool for the extraction and detection of colored molecules from the fermentation broth of filamentous fungi [49]. Among the IL-based systems investigated here, the [N4444]Br⁻-based system appeared to have the best ability to extract fungal pigments, with putative higher affinity for red-orange azaphilone-like pigments. Thus, the IL-ATPS consisting of 15% [N4444]Br⁻ and 25% Na₂CH₃CO₂ was found to be a promising solvent that is worth further investigation to expand its potential applications. Indeed, at this point, the most appropriate use of this system would be for the detection of red azaphilone-like pigments. More studies and optimizations, especially at the back-extraction level, are needed to use this [N4444]Br⁻ system as a powerful extraction technique. Thus, further studies would have to be carried out at larger scales in order to evaluate the obstacles and challenges associated with efficient target molecule recoveries and ionic liquids' recyclability.
**Supplementary Materials:** The following are available online at http://www.mdpi.com/2309-608X/6/4/375/s1, Figure S1: Visual aspect of the reverse extraction steps for each different IL-ATPS tested before (pH = 5) and after adjusting the pH (pH = 13), Figure S2: Back extraction efficiency (A) and overall efficiency (B) of red pigments extraction for each strain in the 4 IL-ATPS tested (n = 3).

**Author Contributions:** Conceptualization, Y.C.; formal analysis, J.L.; funding acquisition, M.F.; investigation, J.L. and Y.C.; methodology, J.L.; project administration, Y.C.; supervision, Y.C.; writing—original draft preparation, J.L.; writing—review and editing, J.L., T.P., M.F., L.D. and Y.C. All authors made critical revisions. All authors have read and agreed to the published version of the manuscript.

**Funding:** This research was funded by Conseil Régional de La Réunion (Réunion Island, France), grant No. DRED 20140704 (COLORMAR Program).

**Conflicts of Interest:** The authors declare no conflict of interest.

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Article

# Microbial Colorants Production in Stirred-Tank Bioreactor and Their Incorporation in an Alternative Food Packaging Biomaterial

Fernanda de Oliveira<fnref n="1" />, Caio de Azevedo Lima<fnref n="1" />, André Moreni Lopes<fnref n="2" />,<fnref n="5" />,⁶

Daniela de Araújo Viana Marques<fnref n="3" />, Janice Izabel Druzian<fnref n="4" />, Adalberto Pessoa Júnior<fnref n="5" /> and

Valéria Carvalho Santos-Ebinuma<fnref n="1" />,*

Received: 28 August 2020; Accepted: 15 October 2020; Published: 2 November 2020

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**Abstract:** Natural colorants from microbial fermentation have gained significant attention in the market to replace the synthetic ones. *Talaromyces* spp. produce yellow-orange-red colorants, appearing as a potential microorganism to be used for this purpose. In this work, the production of natural colorants by *T. amestolkiae* in a stirred-tank bioreactor is studied, followed by its application as additives in bio-based films. The effect of the pH-shift control strategy from 4.5 to 8.0 after 96 h of cultivation is evaluated at 500 rpm, resulting in an improvement of natural colorant production, with this increase being more significant for the orange and red ones, both close to 4-fold. Next, the fermented broth containing the colorants is applied to the preparation of cassava starch-based films in order to incorporate functional activity in biodegradable films for food packaging. The presence of fermented broth did not affect the water activity and total solids of biodegradable films as compared with the standard one. In the end, the films are used to pack butter samples (for 45 days) showing excellent results regarding antioxidant activity. It is demonstrated that the presence of natural colorants is obtained by a biotechnology process, which can provide protection against oxidative action, as well as be a functional food additive in food packing biomaterials.

**Keywords:** natural colorants; filamentous fungi; stirred-tank bioreactor; biodegradable films; antioxidant; food package

## 1. Introduction

Over the years, humans have used colorants to enhance or restore the appearance of products, especially in the food industry [1,2]. Nevertheless, most of the synthetic colorants used in food products are derived from petroleum and may have harmful effects on human health and the environment [3,4]. Therefore, there is a growing preference for natural additives in foods. Regarding the natural color market, a survey by the Grand View Research, Inc. estimated the world color market for 2025 to be

<footnote n="1">¹ Department of Engineering Bioprocess and Biotechnology, School of Pharmaceutical Sciences, Universidade Estadual Paulista—UNESP, Araraquara 14800-903, Brazil; fernanda.oliveira1@unesp.br (F.d.O.); caio.a.lima@unesp.br (C.d.A.L.)</footnote>

<footnote n="2">² Faculty of Pharmaceutical Sciences, University of Campinas—FCF/UNICAMP, Campinas 13083-859, Brazil; amorenilopes@gmail.com</footnote>

<footnote n="3">³ Laboratory of Biotechnology Applied to Infectious and Parasitic Diseases, Biological Science Institute, University of Pernambuco-ICB/UPE, Recife 50100-130, Brazil; daniela_viana@yahoo.com.br</footnote>

<footnote n="4">⁴ Department of Bromatological Analysis, Faculty of Pharmacy, Postgraduate Program in Science of Food, Federal University of Bahia, Salvador 40170-115, Brazil; janicedruzian@hotmail.com</footnote>

<footnote n="5">⁵ Department of Biochemical and Pharmaceutical Technology, University of São Paulo, São Paulo 05508-000, Brazil; pessooajr@usp.br</footnote>

<footnote>* Correspondence: valeria.ebinuma@unesp.br; Tel.: +55-16-3301-4647</footnote>
USD 37.49 billion. Specifically, for the food market, the natural colorant consumption will register a 5.9% compound annual growth rate (CAGR) in terms of revenue, in relation to the period from 2018 to 2025 [5].

There are several biological sources that produce natural colorants, such as insects, vegetables, and microorganisms [2,6]. Colorants from microbial origin, especially fungal ones, are of great interest mainly because of the recent improvements in biotechnology and bioprocessing technologies as well as a full control of the production conditions [2,7,8]. Fungus of the genus *Monascus* stands out for presenting good levels of colorant production. However, *Monascus* species co-produce citrinin, a mycotoxin [9]. Thus, other strains such as *Aspergillus*, *Trichoderma*, *Fusarium*, and *Talaromyces* (formerly *Penicillium* spp.) have been the subject of studies in order to produce colorants [8,10–12]. In this context, *Talaromyces amestolkiae* (former *Penicillium purpurogenum*) has aroused interest in the production potential of stable and non-toxic yellow, orange, and red colorants [10,13–15].

One of the main challenges of using filamentous fungi, such as *Talaromyces*, for metabolite production is the control of microorganism growth and form besides the mass transfer inside the bioreactor [16]. Hyphal concentration greatly influences the fermentation broth viscosity [15]. Meanwhile, the oxygen transfer in submerged cultures is an important factor for efficient fungal growth and an important parameter for a high yield of fungi colorants [17]. At a high cell concentration, the culture medium viscosity increases and it limits the oxygen supply for the cells [4,18]. Increasing the stirring speed can maintain dissolved oxygen concentration at high levels, however, potential damage to the fungal hyphae can limit the impeller speed and, consequently, the oxygen and nutrient transfer capability of a bioreactor [19,20]. In order to control cell growth and colorant biosynthesis, the well-directed process parameter shift represents a valuable control strategy [9,21]. In the production of *Monascus* colorants, the pH-shift strategy has been proven to be efficient in colorants biosynthesis [17,22]. Hence, great efforts to improve the bioreactor cultivation conditions are necessary for enhancing the large-scale production of *Talaromyces* colorants.

Biosynthetically, many fungi colorants are polyketides. Polyketide colorants range in structure from tetraketides to octaketides, which have four or eight C₂ units that contribute to the polyketide chain. The structure of the polyketides does not exhibit localized negatively-charged ions. These molecules often have a polyunsaturated function, i.e., a ring system, one or more carbonyl groups, carbolic acid, and functional ester or amide groups that absorb in the UV-visible spectrum [23]. One class of polyketide colorants are azaphilones, which are hexaketide colorants with pyrone-quinone structures and a chiral quaternary center [24]. Investigations focus on six major azaphilone colorants: Rubropunctamine and Monascorubramine (red); Rubropunctatin and Monascorubrin (orange); Monascin and Ankaflavin (yellow) [25]. One of the most interesting characteristics of azaphilone colorants is their potential use as a functional food colorant, because they may exert antioxidant and antimicrobial activities, relatively low cytotoxicity, as well as immunosuppressive, antiviral, anticancer, and cholesterol-reducing properties [26]. Moreover, our research group previously reported that *T. amestolkiae*-fermented broth presented low cytotoxicity against fibroblast cells and effective antimicrobial activity against *Staphylococcus aureus*, a representative food contaminant [13]. In this sense, bio-based materials with antioxidant properties are gaining attention in the food sector [27], mainly because non-ecofriendly packages are considered an environmental problem [28].

The application of natural compounds in active packaging has been increasingly applied in order to produce materials that interact with packaged foods, positively modifying their sensory and nutritional properties, preventing the deterioration of food, mainly caused by lipid oxidation and microbial growth [29]. Particularly, foods with high-fat content and especially those with a high degree of unsaturation are susceptible to deterioration by oxidation. In this way, alternative packaging technologies based on the inclusion of antioxidants compounds can create a barrier, improving the stability of packaged products sensitive to oxidation [30]. Therefore, the use of the *T. amestolkiae* natural colorants as an added component in bio-based films represents an alternative for increasing the properties of food packaging.
This work addresses the production of natural colorants by *T. amestolkiae* in a stirred-tank bioreactor. The effects of stirring speed and pH-shift as a control strategy on colorant accumulation are evaluated. After the selection of the best operational condition, the fermented broth containing the colorant is incorporated into cassava starch biodegradable films. The bio-based films prepared are characterized with respect to thickness, water activity, and total solid content. In the end, the cassava starch-based films are used to pack butter, supporting the applicability of natural colorants as additives in bio-based films. The effectiveness of the *T. amestolkiae* colorants protection against oxidative action is addressed on the basis of the peroxide content under accelerated oxidative conditions.

## 2. Materials and Methods

### 2.1. Materials

Sucrose and yeast extract were purchased from Synth (São Paulo, Brazil) and Acumedia (Lansing, MI, USA), respectively. Cassava starch (composed by 23.5% amylose and 64.2% amylopectin) was donated by Cargill Agrícola S.A. (Porto Ferreira, SP, Brazil). Commercial butter was obtained from Imperial (Bahia, Brazil). Low-density polyethylene (LDPE) film (0.020 mm thickness and $15.86 \times 10^{-8} \text{ g H}_2\text{O} \cdot \text{mm/m}^2 \cdot \text{h} \cdot \text{kPa}$ water vapor permeability) was purchased from local markets (Salvador, BA, Brazil). All of the other reagents were of analytical grade.

### 2.2. Microorganism Maintenance and Colorant Production

*Talaromyces amestolkiae* DPUA 1275 was generously provided by the Culture Collection of the Federal University of Amazonas (DPUA, Manaus, AM, Brazil). The cultures preserved in distilled water were reactivated in Czapeck Yeast Extract Agar (CYA) and maintained at 30 °C for 7 days. The CYA medium had the following composition (g/L in deionized water): $K_2HPO_4$ (1.0), yeast extract (5.0), sucrose (30.0), agar (15.0), and 10 mL/L of concentrated Czapeck [(g/100 mL of deionized water): $NaNO_3$ (30.0), KCl (5.0), $MgSO_4 \cdot 7H_2O$ (5.0), and $FeSO_4 \cdot 7H_2O$ (0.1)] [10].

The production process was composed of three phases: pre-inoculum, inoculum, and submerged culture. For the pre-inoculum preparation, a loop of fungus from stock culture was inoculated on a CYA plate and maintained in the same reactivation conditions. For inoculum, five mycelial agar discs (8 mm diameter) of *T. amestolkiae* were punched out from the pre-inoculum with a self-designed cutter and transferred to 50 mL of submerged culture medium in 250-mL Erlenmeyer flasks incubated at 30 °C and 150 rpm for 72 h. The inoculum medium was CYA liquid (without the addition of agar). Then, the entire volume obtained (0.2 L) was aseptically transferred to a single flask and then transferred to the bioreactor. The composition of the submerged culture medium was similar to that used for inoculum preparation, except for the concentration of sucrose and yeast extract, which were 48.50 and 11.80 g/L, respectively [10]. Both the inoculum medium and the culture broth had their pH adjusted to 4.5 with HCl (5 M) and were autoclaved at 121 °C for 15 min. The submerged culture was performed in a stirred-tank bioreactor Bioflo® 115 (New Brunswick, Edison, NJ, USA) with 3 L of working volume equipped with two Rushton impellers submersed into the bulk liquid. After sterilization, the bioreactor containing 1.8 L of culture medium was supplied with 0.2 L of the inoculum. The following operational conditions were kept constant by the bioreactor controllers: temperature at 30 °C and aeration rate at 2.0 vvm. The experiments were carried out for 240 h, and every 24 h, an aliquot was withdrawn. At the end of the bioprocess, the fermented broth was filtered first using filter paper Whatman #1 (Whatman, Marlborough, England) and later using a 0.45 µm filter acquired from Millipore. The filtrated samples were used to determine the production of yellow, orange, and red colorants as well as sucrose consumption. In fact, during the cultivation of *Talaromyces* spp., the three natural colorants (yellow, orange, and red) were produced at the same time. In order to guarantee that the colorants were produced, we analyzed the specific wavelength for each one of them, according to Section 2.3.
In the first set of experiments, the effects of the stirring speed from 100 to 600 rpm were evaluated. Next, pH-shift experiments were carried out at 500 rpm, adjusting the pH to 8.0 with NaOH (1 M) after the sucrose depletion (96 h of cultivation).

## 2.3. Analytical Methods

The sucrose concentration was determined according to the methodology described by Dubois et al. [31] and the pH was measured using a pH-meter (model MPC 227, Mettler Toledo, Columbus, OH, USA). The production of extracellular colorants was estimated by spectrophotometric analysis (model DU 640, Beckman, Irving, TX, USA) by reading the absorbance of supernatant at 410, 470, and 490 nm, which corresponds to the maximum absorbance for yellow, orange, and red colorants, respectively. The results were expressed in terms of absorbance units (AU) [10].

## 2.4. Kinetic Parameters

To calculate the substrate conversion factor (sucrose) in product ($Y_{P/S}$) and productivity ($P$), we employed Equations (1) and (2), respectively:

$$Y_{P/S} = \frac{Abs_f - Abs_0}{S_0 - S_f} \qquad (1)$$

$$P = \frac{Abs_f}{t} \qquad (2)$$

where: $S_0$ and $S_f$ are the initial and final sucrose concentrations; $Abs_f$ and $Abs_0$ are the final and initial colorant absorbance, respectively, and $t$ is the time (h).

## 2.5. Preparation of Cassava Starch-Based Films and Incorporation of Natural Colorants

The cassava starch-based films were prepared according to the methodology described by Silva et al. [32]. In this way, the bio-based films were prepared by the casting technique, mixing cassava starch (4.0 wt%), plasticizers (0.7 wt% sucrose and 1.4 wt% inverted sugar), and the fermented broth containing the colorants (5.0 wt%). The fermented broth was used because of the synergy between the colorants molecules and the different metabolites present on fermented broth that increase the antioxidant power. Dispersions were heated (70 ± 2 °C), degassed for 30 min in ultrasonic bath to remove the bubbles, placed in polystyrene Petri dishes (150 × 15 mm), and dehydrated in an oven with airflow and circulation at 35 ± 2 °C for 24 h. A cassava starch-based film without addition of the fermented broth was prepared and used as control. Resulting bio-based films were stored in desiccators (at 23 ± 2 °C with 60 ± 2% relativity humidity) with a supersaturated solution of magnesium nitrate for 48 h before the characterization studies.

## 2.6. Cassava Starch-Based Films Characterization

The thickness was evaluated using a flat parallel surface micrometer (Mitutoyo model 103–137, precision 0.002 mm). Six measurements were taken at random positions around the film sample. The results were expressed as the mean ± respective standard deviations and used for the calculation of the contact area (mm).

The water activity (aw) was measured with a Decagon, Aqualab Lite, as calibration standards used pure water (aw of 1.000 ± 0.001%) and LiCl (aw of 0.500 ± 0.015%). Preconditioned samples (4 cm²) were cut from the center of the films and evaluated in triplicate at room temperature (25 ± 2 °C).

The total solid content was determined by measuring the weight loss of films upon drying (105 °C) until constant weight. This determination was performed in triplicate. The results were expressed as the mean ± respective standard deviations.
## 2.7. Application of Cassava Starch-Based Films for Packaging Butter

Butter was packed in the cassava starch-based films with colorant additives. Square-shaped films (5 × 2 cm) of 0.164 and 0.212 mm in thickness were molded (Sealer Sulpack SM 400 TE, Caxias do Sul, Rio Grande do Sul, Brazil). Butter samples were homogenized and frozen in small pieces (with 3 × 2 cm and 10.00 ± 0.54 g), after they were involved with the film, the bubbles of oxygen were removed, and the film was sealed.

The samples of packaged butter were stored for different periods of time (0, 7, 15, 30, and 45 days) and under accelerated oxidative conditions (64% of relative humidity at 25 ± 2 °C). These analyses were carried out in a dark room to avoid the effects of light interference (i.e., color degradation) in the samples. For this, three types of packages were prepared: cassava starch-based films with colorant additives (CFC), cassava starch-based films without colorant additives (CF), and conventional plastic (CP). In addition, unpackaged butter (U) was used as control. To evaluate the antioxidant action of the films formulation, the oxidative stability of the butter stored under accelerated oxidative conditions was monitored for 45 days and the peroxide content (PC) was determined by titration method according to the methodology described by the Association of Official Analytical Chemists–AOAC (2000) [33].

## 2.8. Statistical Analysis

The data were analyzed by ANOVA using a StatSoft v.7 program (StatSoft, Inc., Tulsa, OK, USA). The Tukey test was used to evaluate the mean difference in results at the 95% level of significance.

# 3. Results and Discussion

## 3.1. Production of Natural Colorants in Bioreactor: Effect of Stirring Speed

The high viscosity of the medium can result in oxygen diffusion limitation, due to the combination of the high biomass concentration and the fungal morphology. Hence, efforts have been made in order to overcome mass transfer problems [16,34]. Therefore, the effects of the stirring speed and pH were investigated by our group. In this sense, the production of natural colorants by *T. amestolkiae* in a stirred-tank bioreactor was evaluated, we maintained the aeration rate constant at 2.0 vvm and varied the stirring speed from 100 to 600 rpm in order to achieve proper oxygen diffusion in our cultivations. The results are depicted in Figure 1.

As can be seen in Figure 1, the stirring speed influenced the colorants' yield. At 100 rpm, the colorants' production stopped after 120 h, probably because under this condition the microorganism distribution inside the bioreactor was not homogeneous. It is usually observed that submerged cultures of filamentous fungi present a pseudo plastic behavior (i.e., a non-Newtonian characteristic) [35], and low speed, associated with high medium viscosity, does not allow for system homogeneity. The main role of stirring in a bioreactor is to improve heat and mass transfer to achieve homogeneity in the system. A proper mixing of components in the culture medium is necessary to ensure an adequate flow in submerged aerobic culture [36]. The stirring of the culture medium may also cause different effects in filamentous microorganisms. Among them may be included: cell wall disruption, changes in filamentous morphology, variation in growth efficiency and growth rate, and the variation in the formation rate of the desired bioproduct [18,36]. The stirring process may damage both mycelial pellets and hyphae structures. The fragmentation of hyphae can result in the creation of small parts of these structures, promoting a new center for biomass growth, or may cause damage to the hypha allowing release of the cytoplasm. On the other hand, total pellet breakdown may occur, and, probably in this case, total autolysis or aggregates can occur [37].
<figure><img src="image_103.png" /><figcaption>**Figure 1.** Production of yellow (yellow bars), orange (orange bars), and red colorants (red bars) by submerged culture of *T. amestolkiae* in stirred-tank bioreactor at 2.0 vvm and 30 °C by varying the stirring speed (100 to 600 rpm) as a function of time (0 to 240 h). The error bars represent 95% confidence levels for the mean of three independent assays.</figcaption></figure>

Increasing the stirring speed could maintain dissolved oxygen concentration at high levels being a key parameter for high yield of *T. amestolkiae* colorants in the stirred-tank bioreactor cultivation. As the stirring speed increased, the homogeneity of the culture medium could be achieved, and it was possible to determine the natural colorant production. Since the oxygen transfer is directly proportional to the shear force and it is strictly related to the morphology of the filamentous fungus, stirring speeds above 600 rpm were not evaluated in this study. Considering the production of orange and red colorants, there was no statistical difference between the results achieved at 500 rpm and 600 rpm (statistical test: one-way ANOVA, *p* = 0.0994). However, as 500 rpm promote a lower shear stress, this stirring speed was chosen for the next experiments. In this condition, the absorbance of the yellow, orange, and red colorants was 1.71 AU<sub>400nm</sub>, 0.71 AU<sub>470nm</sub>, and 0.69 AU<sub>490nm</sub>, respectively. Comparing the colorant production achieved with results previously reported by our research group [10], using the same culture media but in orbital shaker incubator, there was a decrease of 5-fold for the red colorant production in the bioreactor. This result confirms the statement that the scale-up from orbital shaker incubator to large fermenters is a difficult task. In fact, several conditions of cultivation can change in the bioreactor, mainly the oxygen transfer rate in the medium.

Figure 2 shows the results of sucrose consumption and pH over time of submerged culture of *T. amestolkiae* in bioreactor varying the stirring speed. All values are detailed in Table S1 in the supplementary material.
<figure><img src="image_104.png" /><figcaption>**Figure 2.** Consumption of sucrose (A) and pH (B) over time (0 to 240 h) of submerged culture of *T. amestolkiae* in stirred-tank bioreactor varying the stirring speed (100 to 600 rpm) at 2.0 vvm and 30 °C. The error bars represent 95% confidence levels for the measurements.</figcaption></figure>

Regardless of the stirring speed evaluated, the sucrose consumption showed a similar profile (Figure 2A). In general, the concentration of sucrose decreased during the cultivation and a pronounced consumption of sucrose at 96 h occurred. At the end of cultivations, the final sucrose concentration was close to 2 g/L in all conditions studied.

With respect to pH variation, regardless of the stirring speed evaluated, there was an increase in its value throughout the cultivation, with the pH value of 4.5 reaching values close to 7.5 after 240 h of cultivation (Figure 2B). This profile was similar to what occurred in the submerged culture of *T. amestolkiae* (former *Penicillium purpurogenum*) in the orbital shaker incubator studied by Santos-Ebinuma et al. [10]. Therefore, pH 4.5 is suitable to provide good cell growth.

## 3.2. *Production of Natural Colorants by Submerged Culture in Bioreactor: pH-Shift Strategy*

The stirring speed of 500 rpm was selected to evaluate the pH-shift control strategy on *T. amestolkiae* colorant accumulation. At 500 rpm, the initial pH of 4.5 changed to approximately 8.0 at the end of cultivation (Figure 2B). According to Orozco and Kilikian (2008) [38], the production of red colorants by the submerged culture of *Monascus purpureus* in a bioreactor can be favored by a pH change. These authors selected a pH of 5.5 for the growth step and pH 8.0 for the production step. Thus,
a similar strategy was used in this work in order to increase the colorant production in the bioreactor cultivation. In this way, at 500 rpm of stirring speed, a cultivation was carried out and, after 96 h, the pH was changed to 8.0 by the addition of 5 M NaOH. The time of 96 h was selected according to the experiments carried out that varied the stirring speed, because it was observed that, at this point, there was a marked decrease in the concentration of the primary carbon source (Figure 2B). Figure 3 depicts the production of yellow, orange, and red colorants, as well as the pH for the assays performed at 2.0 vvm, 30 °C, and 500 rpm.

<figure><img src="image_105.png" /><figcaption>**Figure 3.** Production of yellow (yellow bars), orange (orange bars), and red colorants (red bars), pH (circle) and sucrose (square) as a function of time (0 to 240 h) during the submerged culture of *T. amestolkiae* in a stirred-tank bioreactor at 2.0 vvm, 30 °C, and 500 rpm. The black arrow indicates the pH-shift from 4.5 to 8.0 at 96 h for the production step. The error bars represent 95% confidence levels for the measurements.</figcaption></figure>

After the pH-shift, it is possible to observe a significant increase in the production of all the colorants, with this increase being more significant for the orange and red ones, both close to 4-fold. Furthermore, the influence of the pH-shift on colorant production was evident. The maximum yield of yellow, orange, and red colorants was 3.20 AU<sub>400nm</sub>, 2.56 AU<sub>470nm</sub>, and 2.45 AU<sub>490nm</sub>, respectively. These results corroborate those found by Orozco and Kilikian (2008) [38], who mention that different pH levels during the growth and production steps may lead to an improvement in the colorant production. Although the colorant production with the pH-shift strategy was greater than for the first set of experiments, the product yield was still lower than for the best condition in the orbital shaker incubator, which shows that further studies are necessary to improve the production of *T. amestolkiae* natural colorants in a stirred-tank bioreactor.

Compared with no pH change condition, all kinetic parameters evaluated improved with the pH-shift control strategy (Table 1). The rate of the substrate conversion to product increased from 0.037 to 0.067 AU·L/g for the yellow colorants and from 0.015 to 0.054 and 0.051 AU·L/g for the orange and red colorants, respectively. Similarly, the productivity of all colorants increased in this second experimental condition.

Generally, in bioreactor cultivation, reproducible kinetic growth of filamentous organisms is difficult to obtain. This phenomenon occurs frequently, due to mycelial aggregation in fermentation broths with long hairy mycelial morphologies that can adhere to surfaces and form a growing biofilm [39]. Many fungal strains grow preferentially on surfaces and will develop thick layers on walls and surfaces in a submerged fermentation process [40]. In the same way, reproducible samples to measure apparent viscosity are difficult to obtain, hence, it was also deemed appropriate to omit the time course for biomass and viscosity.
<table><caption>**Table 1.** Kinetic parameters calculated for substrate conversion factor (sucrose) in product (Y<sub>P/S</sub>) and productivity during the submerged cultivation of *T. amestolkiae* in a stirred-tank bioreactor for the production of yellow (Y, AU<sub>400nm</sub>), orange (O, AU<sub>470nm</sub>), and red colorants (R, AU<sub>490nm</sub>) without a pH change and after a pH-shift from 4.5 to 8.0 at 96 h of cultivation.</caption><thead><tr><th rowspan="2">Experimental Condition</th><th colspan="3">Y<sub>P/S</sub> (AU·L/g)</th><th colspan="3">Productivity (AU/h)</th><th rowspan="2">Abs<sub>O</sub>/Abs<sub>Y</sub></th><th rowspan="2">Abs<sub>O</sub>/Abs<sub>R</sub></th></tr><tr><th>Y</th><th>O</th><th>R</th><th>Y</th><th>O</th><th>R</th></tr></thead><tbody><tr><td>Without pH change</td><td>0.037</td><td>0.015</td><td>0.015</td><td>0.007</td><td>0.007</td><td>0.007</td><td>0.415</td><td>1.025</td></tr><tr><td>pH-shift strategy</td><td>0.067</td><td>0.054</td><td>0.051</td><td>0.013</td><td>0.011</td><td>0.010</td><td>0.800</td><td>1.049</td></tr></tbody></table>

Abs<sub>O</sub>/Abs<sub>Y</sub>: relationship between the production of orange and yellow colorants; Abs<sub>O</sub>/Abs<sub>R</sub>: relationship between the production of orange and red colorants.

Even though it is desirable to produce fungal colorants comprised of only one single color component [22], it is known that the fermented broth is composed of yellow, orange, and red colorants [13–15]. However, the careful selection of pH and nitrogen sources is a valid approach to produce extracellular extracts comprised of one predominant color component [14]. It has been widely accepted by the scientific community that orange colorants are the first biosynthetic product and the other colorants derive from the orange ones [41]. In this way, the extracellular colorant composition and concentration is dependent of the orange colorants produced that are transformed into red and yellow colorants under some specific fermentation conditions. For the *Monascus* genus, a low pH (pH 2.5 and 4.0) provides intracellular extracts composed mainly of orange colorants, independently of the nitrogen source employed. At acidic conditions, the secretion of orange colorants into the broth and the reaction with any amino unit for red colorants formation is limited [22]. Meanwhile, a change of pH to levels closer to neutral modify the extent of transformation of orange colorants into red ones [22]. Additionally, for *T. amestolkiae*, the synergistic effect of a low pH and nitrogen source is mainly important in the production and excretion of red colorants. A previous study of our research group presented the production of a glutamic acid-red colorant complex by *T. amestolkiae* in a chemically defined medium with monosodium glutamate (MSG) as the nitrogen source [14]. In the presence of MSG, deep yellow colorants were derived from neutral and basic pH, while deep red colors were derived from acidic pH. The glutamic acid–colorant complex seems to be more water-soluble than those produced in nitrogen complex media (meat extract and meat peptone) by the same strain [13].

In this work, yeast extract was used as complex nitrogen source and the synergistic effect of this nitrogen source with pH favored the increase of the relation between orange colorants and yellow/red ones. This is a reflection of the overall increased rate of substrate to product and productivity. It seems that a close to neutral pH favored the reduction of orange colorants to yellow ones (relation Abs<sub>O</sub>/Abs<sub>Y</sub> < 1.0 duplicated with pH-shifting), while the specific red conversion was limited (relation Abs<sub>O</sub>/Abs<sub>R</sub> > 1.0). Consequently, the extracellular extracts contained mainly yellow colorants. Therefore, extracellular red colorants derivatives were not mainly produced at a high pH. It can be considered that natural colorants can be obtained by the submerged culture of *T. amestolkiae* in a bioreactor, however, the culture conditions can be improved and the production increased by varying mainly the pH during cultivation. In order to study a possible application of the colorants produced, the incorporation of the fermented broth into an alternative food-packaging material based on its oxidative protection effect was evaluated.

### 3.3. Preparation and Characterization of Cassava Starch-Based Films

Biodegradable films were produced by a formulation of cassava starch, plasticizers, and the fermented broth containing the *T. amestolkiae* colorants produced in stirred-tank bioreactor. The incorporation of natural additives from plants for active packaging has been increasingly applied, such as: coffee-cocoa [42,43], carotenoid and yerba mate extract [44], green tea and palm oil extract [45],
among others. However, natural additives produced by biotechnology is an innovative approach. The bio-based films produced were characterized in terms of thickness, water activity, and total solids content. The results are depicted in Table 2.

<table><caption>**Table 2.** Characterization of biodegradable films in the presence or absence of natural colorants in terms of thickness, water activity (aw), and total solids.</caption><thead><tr><th>Characterization/Parameters<br>Analyzed</th><th>Biodegradable Films<br>Containing Natural Colorants *</th><th>Biodegradable Films Without<br>Natural Colorants *</th></tr></thead><tbody><tr><td>Thickness (mm)</td><td>0.147 ± 0.009 <sup>a</sup></td><td>0.163 ± 0.013 <sup>b</sup></td></tr><tr><td>aw (%)</td><td>0.612 ± 0.041 <sup>a</sup></td><td>0.604 ± 0.050 <sup>a</sup></td></tr><tr><td>Total solids</td><td>89.80 ± 0.540 <sup>a</sup></td><td>88.96 ± 0.500 <sup>a</sup></td></tr></tbody></table>

The physical properties of the films were determined to evaluate the influence of the colorant incorporation, since they could modify the structure of the polymer matrix, e.g., by weakening the inter-chain bonds. The incorporation of natural colorants in the film formulation caused a significant reduction of film thickness, around 10%, in relation to films with no additives. Other works also reported a reduction in thickness of films formulated with cassava starch and natural additives [43,46]. The control of the thickness in the biodegradable films is an important step, since variations in this parameter can affect some films properties, such as mechanical and barrier properties, which can compromise the package performance [45]. Thickness was also the only parameter addressed that was affected significantly.

The variations in the total solid content in the films after colorant incorporation were not significant, with values of 88.96 and 89.80%, respectively. In general, the incorporation of natural additives in cassava starch-based films does not affect significantly the solid content [43].

Additionally, no significant difference (<0.05) for water activity (aw) between the films produced was observed. These results demonstrate that the presence of natural colorants did not change the water activity of the films. Water activity brings information about the amount of free (also referred as unbound or active) water present in a sample [47] being a parameter used in food preservation. Low water activity reduces the availability of water to microorganisms, avoiding undesirable chemical changes for the storage of products [32]. As water migrates from areas of high aw to areas of low aw, it is important to have low values of aw (<0.600 according to Mathlouthi, 2001 [48]) for food product design. As the aw was around 0.6 for both films, they can be considered as food product packages. In this way, the cassava starch-based films were used to pack butter samples.

### 3.4. Oxidative Stability of the Packaged Butter during Storage

The oxidative stability of the butter packaged in different formulations, namely: cassava starch-based films with colorant additives (CFC), cassava starch-based films without colorant additives (CF), and conventional plastic (CP) was evaluated. Unpacked butter (U) was used as the control. To this purpose, the packed or unpackaged butter was stored for 45 days under accelerated oxidative conditions (64% of relative humidity and 25 °C) and monitored through peroxide content (PC) at 7, 15, 30, and 45 days of storage. Figure S1 from Supplementary Material shows an image of the butter packaged in each formulation evaluated. Figure 4 shows the results of PC for each condition over the 45 days of storage. All values are detailed in Table S2 in Supplementary Material.

<footnote>* Statistical test: one-way ANOVA (with Welch correction for unequal variances). The results were expressed as the mean ± respective standard deviations. Equal letters in the same line represent equal values to a significance level of 5%.</footnote>
<figure><img src="image_106.png" /><figcaption>**Figure 4.** Peroxide content (PC, meq/kg) in the butter as a function of the packaging film used, such as: cassava starch-based films with colorant additives (CFC), cassava starch-based films without colorant additives (CF), conventional plastic (CP), and unpackaged butter (U), during different periods of time (0, 7, 15, 30, and 45 days) in storage. The error bars represent 95% confidence levels for the measurements.</figcaption></figure>

From Figure 4 can be seen that, as expected, the PC of the U was higher than of the packaged one. The CF and CP showed higher peroxide indices than CFC. This demonstrates that natural colorants act as both a photooxidative protector and antioxidant agent, which allow a higher oxidative protection of the packaged product when compared to the other conditions evaluated.

After 45 days of storage, the PC increased 329.0% for the exposed butter, 239.7% for the CP, and 231.1% for the CF. While for the CFC, the PC increased only 118.5%. Therefore, even without antioxidant additives (CF), cassava starch-based films have a greater oxidative protection effect when compared to conventional package films (CP). However, the formulation with colorants incorporated in cassava starch-based films (CFC) enhanced the oxidative protection of butter, with a reduction in peroxide value. This shows a synergistic action of the bio-based film and the colorants, improving the shelf life of fatty products. In this way, *T. amestolkiae* colorants can be considered as potential natural antioxidants for the stabilization of lipid-containing foods for active packing formulation. However, additional studies on the concentration of colorants in the formulation should be performed in order to avoid a possible pro-oxidant effect. Since a high content of antioxidant additives can act as a pro-oxidant agent [44,45].

**4. Conclusions**

This work evaluated the influence of the stirring speed and pH on the production of natural colorants by *T. amestolkiae* in a stirred-tank bioreactor. The greatest colorant production occurred at 500 rpm, under the pH-shift strategy from 4.5 to 8.0 during the production phase. Although the regulatory mechanisms of the biosynthesis of *T. amestolkiae* colorants are not yet clear, the relationship between the culture conditions and the colorant formation it is of great importance. Despite the difficulties in terms of viscosity of the medium, during the submerged cultivation of filamentous fungi in a bioreactor, there is the possibility of using strategies to control growth and metabolite production, especially in terms of stirring and pH. Moreover, this work demonstrated the potential of natural colorants to be included in biodegradable film formulation. The incorporation of colorants in cassava starch-based films provides oxidative protection in packaged butter, by the decrease of peroxide index. Hence, *T. amestolkiae* colorants have great potential for use as functional food colorants. As future perspectives, our group intends to evaluate the surface morphologies of the films with or
without colorants by scanning electron microscope in order to gather supplementary data about the
characteristics and stability of our formulations extending, for example, its shelf-life application.

**Supplementary Materials:** The following are available online at http://www.mdpi.com/2309-608X/6/4/0264/s1,
Figure S1: Butter packaged in cassava starch-based films with colorant additives (CFC), cassava starch-based films
without colorant additives (CF), conventional plastic (CP), and unpackaged butter (U), Table S1: Consumption of
sucrose and pH over time (0 to 240 h) of submerged culture of *T. amestolkiae* in stirred-tank bioreactor varying the
stirring speed (100 to 600 rpm) at 2.0 vvm and 30 °C, Table S2: Peroxide content in the butter as a function of the
packaging film used, such as: cassava starch-based films with colorant additives (CFC), cassava starch-based films
without colorant additives (CF), conventional plastic (CP), and unpackaged butter (U), during different periods of
time (0, 7, 15, 30, and 45 days) in storage.

**Author Contributions:** Conceptualization, F.d.O., C.d.A.L., V.C.S.-E.; methodology, F.d.O., J.I.D., A.P.J., V.C.S.-E.; software, F.d.O., V.C.S.-E.; validation, D.d.A.V.M., A.P.J., V.C.S.-E.; formal analysis, V.C.S.-E.; investigation, F.d.O., V.C.S.-E.; resources, A.P.J., V.C.S.-E.; data curation, A.P.J., V.C.S.-E.; writing—original draft preparation, F.d.O., C.d.A.L., A.M.L., V.C.S.-E.; writing—review and editing, F.d.O., A.M.L., D.d.A.V.M., V.C.S.-E.; visualization, A.P.J., V.C.S.-E.; supervision, J.I.D., A.P.J., V.C.S.-E.; project administration, V.C.S.-E.; funding acquisition, A.P.J., V.C.S.-E. All authors have read and agreed to the published version of the manuscript.

**Funding:** This work was developed with the financial support from FAPESP (São Paulo Research Foundation, Brazil) through the projects 2018/06908-8, 2014/01580-3, 2013/05039-2, 2008/58280-0. A.M. Lopes acknowledges the financial support from FAPESP through the projects 2017/10789-1 and 2018/10799-0. This study was financed in part by the Coordenação de Aperfeiçoamento de Pessoal de Nível Superior - Brasil (CAPES) - Finance Code 001. The authors also acknowledge the support from the CNPq (National Council for Scientific and Technological Development, Brazil).

**Conflicts of Interest:** The authors declare no conflict of interest.

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Article

# Oil-Based Fungal Pigment from *Scytalidium cuboideum* as a Textile Dye

Mardonio E. Palomino Agurto<fnref n="1" />, Sarath M. Vega Gutierrez ¹ID, R. C. Van Court ¹ID,
Hsiou-Lien Chen <fnref n="2" /> and Seri C. Robinson <fnref n="1" />,*

Received: 31 March 2020; Accepted: 20 April 2020; Published: 22 April 2020

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**Abstract:** Identification of effective natural dyes with the potential for low environmental impact has been a recent focus of the textile industry. Pigments derived from spalting fungi have previously shown promise as textile dyes; however, their use has required numerous organic solvents with human health implications. This research explored the possibility of using linseed oil as a carrier for the pigment from *Scytalidium cuboideum* as a textile dye. Colored linseed oil effectively dyed a range of fabrics, with natural fibers showing better coloration. Scanning electron microscopy (SEM) revealed a pigment film over the fabric surface. While mechanical testing showed no strength loss in treated fabric, colorfastness tests showed significant changes in color in response to laundering and bleach exposure with variable effects across fabric varieties. SEM investigation confirmed differences in pigmented oil layer loss and showed variation in pigment crystal formation between fabric varieties. Heating of the pigmented oil layer was found to result in a bright, shiny fabric surface, which may have potential for naturally weatherproof garments.

**Keywords:** fungal pigment; natural dye; spalting; *Scytalidium cuboideum*; dramada; sustainable clothing

## 1. Introduction

Natural dyes and pigments have a long history of use for coloring textiles, from ancient Egypt to the oldest South American cultures [1–5]. Artificial dyes currently dominate the market due to their ease of mass production, low price, and color variety [5]. However, most modern textile colorants are produced using hazardous chemicals and many contribute to water pollution through the production of effluents [6,7]. In recent decades, the low environmental impact and sustainability of natural dyes have become more desired by consumers, driving a shift in the market back to natural sources of coloration [8,9].

There are presently a number of natural colorant alternatives available, such as those derived from barks, fungal pigments, insects, and minerals [3]. Most have never been commercialized due to concerns about their sustainability, cost effectiveness, and profitability [10]. In addition, natural dyes often have problems with colorfastness, and, while methods have been developed to improve dye uptake [11–13], they are energy intensive and increase the production price [14–16].

One source of natural coloration that has been suggested as a replacement for synthetic dyes in fabric dyeing are pigments extracted from wood-rotting fungi. Research by Weber et al. [17] has shown that fungal pigments, when carried in dichloromethane (DCM), with and without mordants, showed promise for dyeing fabrics, using a dripping process. The pigments from four fungi were investigated: *Chlorociboria aeruginosa* (Oeder), Seaver, and *C. aeruginascens* (Nyl.) Kanouse,

<footnote n="1">¹ Department of Wood Science and Engineering, Oregon State University, Corvallis, OR 97331, USA; mardonio.palomino@oregonstate.edu (M.E.P.A.); sarathth@yahoo.co.uk (S.M.V.G.); ray.vancourt@oregonstate.edu (R.C.V.C.)</footnote>

<footnote n="2">² College of Business, Oregon State University, Corvallis, OR 97331, USA; hsiou-lien.chen@oregonstate.edu</footnote>

<footnote>* Correspondence: seri.robinson@oregonstate.edu; Tel.: +1-541-737-4233</footnote>