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                    Applied Microbiology and Biotechnology (2022) 106:8021–8034
https://doi.org/10.1007/s00253-022-12255-9

BIOTECHNOLOGICAL PRODUCTS AND PROCESS ENGINEERING

Chemical characterization and microencapsulation of extracellular
fungal pigments
Paulina I. Contreras‑Machuca1,2 · Marcia Avello1 · Edgar Pastene3 · Ángela Machuca4 · Mario Aranda5 ·
Vicente Hernández6,7 · Marcos Fernández2
Received: 11 July 2022 / Revised: 18 October 2022 / Accepted: 23 October 2022 / Published online: 12 November 2022
© The Author(s), under exclusive licence to Springer-Verlag GmbH Germany, part of Springer Nature 2022

Abstract
In this work, extracellular colored metabolites obtained from the filamentous fungi Talaromyces australis and Penicillium
murcianum, isolated in the Andean-Patagonian native forests of Chile, were studied as prospect compounds to increase the
sustainability of cosmetic products. The chemical and antioxidant properties of these natural pigments were characterized
and strategies for their microencapsulation were also studied. UHPLC/MS–MS analyses indicated that the predominant
metabolites detected in the cultures of P. murcianum were monascin (m/z = 411.15) and monashexenone (m/z = 319.10),
while athrorosin H (m/z = 458.20) and damnacanthal (m/z = 281.05) were detected in cultures of T. australis. ORAC tests
revealed that P. murcianum’s metabolites had the greatest antioxidant properties with values higher than 2000 μmol of
trolox equivalents/g. The fungal metabolites were successfully microencapsulated by ionic gelation into structures made
of 1.3% sodium alginate, 0.2% chitosan, and 0.07% hyaluronic acid. The microencapsulation process generated structures
of 543.57 ± 0.13 µm of mean diameter (d50) with an efficiency of 30% for P. murcianum, and 329.59 ± 0.15 µm of mean
diameter (d50) and 40% efficiency, for T. australis. The chemical and biological characterization show the biotechnological
potential of these fungal species to obtain pigments with antioxidant activity that could be useful in the cosmetic industry.
The encapsulation process enables the production of easy-to-handle dry powder from the fungal metabolites, which could
be potentially marketed as a functional cosmetic ingredient.
Key points
• The predominant fungal pigments were of azaphilone and anthraquinoid classes.
• The fungal pigments showed high antioxidant activity by ORAC assay.
• Fungal pigment microcapsules obtained by ionic gelation were characterized.
Keywords Fungal pigments · Azaphilones · Anthraquinoids · Mass spectrometry · ORAC assay · Ionic gelation

Introduction
Today, the society exerts greater pressure on products manufactured from novel raw materials, with a reduced content of
additives and synthetic chemicals that can increase toxicity.
This is due to the different risks that can be presented either
during the research and development of new products, or in
its impact once it is distributed in the population. In this way,
* Paulina I. Contreras‑Machuca
paucontrerasm@udec.cl; paucontrerasm@hotmail.com
* Ángela Machuca
angmachu@udec.cl
Extended author information available on the last page of the article

greater environmental regulations have been implemented
in some countries, where the use of many synthetic chemical compounds has been limited or prohibited, encouraging
the search of new molecules and technologies that are more
environmentally friendly and safe for those who manipulate
or use them (Chemat et al. 2012). For this reason, there is a
great interest in natural products derived from microorganisms, since they have the advantage that can be easily cultivated and optimized for the production of certain metabolites such as pigments (Patil et al. 2016; Li et al. 2018).
Natural pigments have traditionally been obtained from
plants, but various problems related with their production
have led to the search for alternative sources such as fungi
(Caro et al. 2015). Fungal pigments, unlike plant pigments,
have the advantage of being mainly extracellular metabolites

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and therefore the recovery from the culture broths is a relatively simple process (Sajid and Akbar 2018). In addition,
the fungal pigments can be produced under laboratory conditions, in confined spaces, regardless of seasonality (Galindo
et al. 2007). Due to this, several filamentous fungi belonging
to the genera Monascus, Fusarium, Penicillium, and Talaromyces, among others, are being investigated as important
sources of natural pigments (Feng et al. 2012; Morales et al.
2020; Pitt and Hocking 2022). Among that, Monascus pigments have been used by humans since ancient times, and
therefore, these pigments are among the most extensively
investigated and described in the literature (Gao et al. 2013;
Gmoser et al. 2017; Chaudhary et al. 2022).
To date, a large number of polyketide compounds have
been described in fungi, of which an interesting group with
multiple beneficial properties corresponds to pigments of
the azaphilone type (Feng et al. 2012; Pimenta et al. 2021).
The Monascus yellow pigment azaphilone type, monascin,
has been related to the reduction of oxidative stress in animal models, activating enzymes and transcription factors
involving in metabolism, resistance to stress, and antioxidant
reserves (Shi and Pan 2012). Another group of polyketide
pigments that are present in species of filamentous fungi of
the genera Penicillium, Talaromyces, and Aspergillus are the
hydroxyanthraquinoids, which also shows important antioxidant and antimicrobial activities (Caro et al. 2012; Li et al.
2017). For these reasons, antioxidant fungal pigments show
a great potential in the formulation of dyes safer than synthetic dyes of traditional use such as azo, triphenylmethane,
and xantene derivatives, widely used in the cosmetic industry, but having a history of cytotoxicity (Crebelli et al. 1981;
Combes and Haveland-Smith 1982). In accordance with the
above, it is necessary to increase the search and characterization of new fungal pigments with biological properties of
interest and potential use in the cosmetic industry (Pimenta
et al. 2021). Penicillium and Talaromyces species are promising because they are neither mycotoxigenic nor pathogenic
for humans, and on the contrary, they can present beneficial
characteristics for health. These include antioxidant, antimicrobial, and anti-inflammatory properties (Gao et al. 2013;
Gmoser et al. 2017; Morales et al. 2020; Poorniammal et al.
2021). Pigments obtained from Talaromyces and Penicillium
species are generally considered of low toxicity. However,
the successful application of these metabolites as new functional ingredient depends on their correct characterization
and the results of toxicity studies performed in accordance
with EU and FDA regulations (Morales et al. 2020; Poorniammal et al. 2021). In specific, toxicity studies with pigments from these fungi revealed a low toxicity and their
suitability to be used in textile applications (Hernández et al.
2019). P. murcianum and T. australis, isolated from decay
wood in forest of central-south Chile, have demonstrated to
be efficient pigment producers. The pigments from these

two fungal species have already been used in the dyeing of
fabrics for textile purposes (Hernández et al. 2018a,b, 2019,
2020), but they have not yet been chemically characterized.
The stabilization of natural pigments for their posterior
incorporation into cosmetic formulation can be achieved by
microencapsulation. This process grants protection and special properties to the encapsulated product. For instances,
encapsulation can provide protection against adverse environments, stabilize sensitive drugs, and generate a controlled
or prolonged release system. Microencapsulation can be
used to protect natural pigments from oxidative processes,
light, humidity, etc., maintaining a more intense and prolonged coloration (Swarbrick 2006; Jurić et al. 2020).
The main objective of the present study was to characterize the extracellular colored pigments produced by the
filamentous fungi Penicillium murcianum and Talaromyces
australis, evaluating their biological activity through antioxidant capacity, and determining the viability of its microencapsulation. Studies on the chemical nature of these fungal
pigments are necessary in order to support their use in new
application, in areas such as the cosmetic industry (Pimenta
et al. 2021).

Material and method
Fungal strains and liquid cultures for pigment
production
The filamentous fungi Penicillium murcianum PM 2015 and
Talaromyces australis TA 2015 were isolated of decayed
wood samples collected in central Chile. The fungi were
identified through molecular techniques, by sequencing of
the ribosomal DNA ITS region (Hernández et al. 2018b),
and deposited under code RGM 2467 (P. murcianum)
and RGM 2468 (T. australis) in the Chilean Collection of
Microbial Genetic Resources (CChRGM) at the Instituto de
Investigaciones Agropecuarias (INIA, Chile). This entity is
associated to the World Data Centre for Microorganisms
(WDCM) under the accession acronym CChRGM and
number WDCM 1067. These fungi were grown in flasks
containing 500 mL of optimized nutrient medium, peptoneglucose-yeast extract (PGY), adjusted at pH 5 for T. australis and pH 9 for P. murcianum (Hernández et al. 2019).
The PGY medium consisted of 12 g/L glucose and 0.1 g/L
yeast extract for both fungi, with different peptone concentrations, 11 g/L for T. australis and 20 g/L for P. murcianum.
In addition, the medium for P. murcianum was supplemented
with NaCl (6 g/L). After a 20-day incubation period, at
28 °C under agitation (120 rpm), the cell-free extracts were
obtain after paper filtration. The filtered extracts containing
extracellular pigments of each species were then frozen at
− 18 °C and kept in these conditions until use.

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Applied Microbiology and Biotechnology (2022) 106:8021–8034

Crude extracts
Initially, a glass column (60 × 10 cm) filled with Amberlite XAD-16 resin (Sigma-Aldrich) was used in order to
separate the extracellular fungal pigments from salts and
compounds highly soluble in water. For that, 100 mL of
each filtered extract was added to the column and once
the color was absorbed by the resin, it was washed several
times with distilled water until the supernatant became
transparent. Then, the adsorbed pigments were eluted
from the column with methanol and the collected fractions
were concentrated in a Heidolph (Schwabach, Germany)
rotary evaporator Laborota 4000/4001 efficient, getting a
yellow-brown crude extract for P. murcianum and a redyellow crude extract for T. australis. The yield of the crude
extracts was expressed as milligram of extract per milliliter of filtered broth.

Separation by centrifugal partition chromatography
(CPC)
Crude extracts were fractionated using the CPC instrument
Armen (Armen, France) Spot-CPC 250-L centrifugal partition chromatograph with a 250 mL total cell volume and
a four-way switching valve that allows operation in either
the descending or ascending modes (Ying et al. 2014).
The CPC system was connected to a SPOT-PREP-II system, with integrated UV detector and fraction collector,
and Armen Glider software package. The separation by
CPC was performed in descending mode with a two-phase
solvent systems composed of ethyl acetate/butanol/water
1:1:2 (v/v/v) for P. murcianum and 4:1:5 (v/v/v) for T.
australis. In the second case, another solvent system with
butanol/water 1:1 (v/v) was developed to improve the separation of the most polar pigments. The solvent mixture
was automatically generated by the SPOT-PREP-II unit.
The CPC rotor was first filled with organic phase at 30 mL/
min and 500 rpm rotation. Lower phase was pumped into
the system in the descending mode at a flow rate of 24 mL/
min and rotation was increased from 0 to 2200 rpm for P.
murcianum and 1800 rpm para T. australis. After equilibrium was reached, the samples were dissolved in 10 mL
1:1 mixture of upper and lower layers and injected into the
CPC system (10 mL sample loop). Elution was monitored
using scan 280, 350, 430, and 510 nm wavelengths, collecting 45 fractions in 20 mL tubes (Figure S1). Fractions
were analyzed by HPLC and those with similar composition were combined in fractions according with their online UV–Vis spectra obtained from the preparative detector. The samples that presented the highest amount of pure
pigment were selected from the chromatographic analysis.

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Analysis of fractions by ultra‑high‑performance
liquid chromatography (UHPLC) with a diode array
detector (DAD) and tandem mass spectrometry (MS
and MS–MS)
Chromatography was performed in Shimadzu (Kyoto, Japan)
Nexera X2 UHPLC system composed of LC-30 AD pump,
DGU-20A5R degassing unit, SIL-30 AC autosampler, CTO20 AC column oven, CBM-20A communication module,
SPM-M20A diode array detector (DAD), and LCMS-8030
triple quadrupole (TQ) mass spectrometer (MS) equipped
with an electrospray ionization source (ESI). MS analysis
was performed using the following settings: ESI (−) voltage of 4.5 kV; nebulizer gas ­(N2) flow: 3.0 L/min; drying
gas ­(N2) flow: 15 L/min; desolvation line (DL) temperature: 250 °C; and heat block temperature: 400 °C. Data were
acquired, recorded, and analyzed by means of the Shimadzu
LabSolution 5.8 software. The methodology was developed
as described for Monascus-type pigments with modifications (Inoue et al. 2010). A Kromasil 100-5-Phenyl column
of 4.6 × 250 mm, particle size 5 μm, series M05PHA25/
E164679, at 40 °C was used. The injection volume was
20 µL for all samples. A mobile phase gradient was established, consisting of 0.2% formic acid in water (solvent A)
and 0.2% formic acid in methanol (solvent B). In this case, it
was started at 0 min, 55% of solvent B, gradually increasing
it to 90% at 15 min and during the following minutes, the
gradient was balanced until reaching the proportions of the
initial phase system at 30 min. Assays were performed with
a flow rate of 0.6 mL/min. The analysis was performed with
electrospray ionization (ESI) in negative polarity and two
types of analysis were performed: scan mode (SM), in this
case the mass values recorded were 50–2000 mass–charge
ratio (m/z), and product ion scan (PIS), where the fractional
m/z were selected, which corresponded to the values of the
main pigments.

Evaluation of antioxidant capacity through assays
of oxygen radical absorption capacity tests (ORAC)
Crude extracts and purified fractions of both fungal species were evaluated by monitoring the fluorescence decay
in a fluorescein target oxidized by the 2,2′-azobis (2-methylpropionamidine) dihydrochloride (AAPH) radical, in a
Fluorescence Spectrometer Perkin Elmer (Waltham, MA,
USA). The ORAC-FL assay was performed according with
literature (Ou et al. 2013), with some modifications. In a
96-well black plate, 150 μL of 0.11 μM fluorescein was
added to each well. Then, 25 μL phosphate buffer (blank)
and Trolox® (standard curve between 25 and 50 µM) were
added to each well. The assay was placed to incubate for
30 min at 37 °C. Then, 25 μL AAPH was quickly added
to each well and measurements were taken up between 0

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and 120 min, measuring every 1 min, with an excitation
and emission wavelength of 485 and 539 nm, respectively
at 37 °C. A calibration curve was obtained through the equation of a linear regression of the area under the fluorescein
decrease curve (AUC), versus the Trolox® concentration.
ORAC values for the samples were obtained using the calibration curve and expressed as µmoles of Trolox® equivalents (TE) per gram of sample (µmol TE/g).

Microencapsulation of fungal pigments by vibration
nozzle
Hybrid microcapsules were made by ionic gelation following the instructions provided on the operating manual of
the B-390 BÜCHI microencapsulator (Flawil, Switzerland)
(BÜCHI Labortechnik AG 2016) and literature (Yan et al.
2015; Athamneh et al. 2019), with some modifications. For
this, T. australis and P. murcianum crude extracts (treated
with Amberlite XAD-16) were added separately in a mixture
of 1.3% w/v sodium alginate and 0.07% w/v hyaluronic acid
(95/5) to a final volume of 100 mL. For P. murcianum, the
parameters of the process used on the microencapsulation
were fixed as follows: vibrational frequency of the membrane: 400 Hz; electrode potential: 2000 V; air pressure:
229 mbar; flow: 15 mL/min; nozzle temperature: 40 °C;
and nozzle diameter: 450 µm. For T. australis, the parameters were as follows: vibrational frequency of the membrane: 700 Hz; electrode potential: 2300 V; air pressure:
462 mbar; flow: 8 mL/min; nozzle temperature: 40 °C; and
nozzle diameter: 300 µm. The microdroplets expelled from
the equipment were received in an ionic bath containing a
mixture of 0.2 M calcium chloride and 0.2% w/v chitosan
(3/1) under constant stirring. The obtained microcapsules
were then vacuum filtered through a 10-μm membrane filter,
washed with 200 mL of nanopure water, frozen at − 20 °C,
and finally dried in a Xiang Yi LGJ-10C (China) lyophilizer
for 24 h.

Color of fungal pigment microcapsules
The color of the fungal pigments microcapsules was measured on a CM-5 Konica Minolta (NJ, USA) spectrophotometer. For the measurements, a 0.60 g sample was taken from
each type of fungal microcapsule and deposited on different
transparent plastic plates. Reflectance reading on the spectrophotometer were then carried out, registering the color
on the CIE L* a* b* coordinates, where L* corresponds to
lightness from 0 (black) to 100 (white), a* from – 60 to + 60
(greenness-blueness), and b* from –60 to + 60 (bluenessyellowness) (Mapari et al. 2006). In addition, from the coordinates, the values c* (saturation) and h* (hue) were also
obtained (Ohta and Robertson 2005).

Granulometric analysis by optical and scanning
electron microscopy
In order to evaluate the microencapsulation process, fungal pigment microcapsules were first observed under a
Primo Star Zeiss (Germany) optical microscope, with a
4× magnification. Later on, a Tescan Vega 3 SBU Easyprobe (Kohoutovice, Czech Republic) scanning electron
microscope (SEM) was used to analyze the microcapsules.
SEM was performed at the Center for Advanced Microscopy, CMA Bio-Bio, Concepción, Chile.
Morphology of microcapsules was defined according
to the classification described in United States Pharmacopeia No. 30 (USP 30) (Convention 2007b) for light
microscopy. For size distribution determination, the Feret
diameter technique described in USP 30 (Convention
2007a) was used. This corresponds to the distance between
imaginary parallel lines tangent to a randomly oriented
particle perpendicular to the eyepiece scale. Observations
were made on samples of 300 microcapsules per batch.
Data were processed to obtain a frequency distribution
table ordered by class intervals, in order to obtain the
geometric and statistical mean diameters (Aulton 2002;
Montenegro et al. 2011). Size distribution of the microcapsules was described quantitatively using the particle
size dispersal coefficient (Span) calculated according to
Span = (d90−d10)/d50, where dn (n = 10, 50, and 90) denotes
the particle diameter at 10%, 50%, and 90% of the size distribution. A low Span is indicative of a more homogeneous
size distribution (Xiao et al. 2005).

Determination of encapsulation efficiency
by spectrophotometry
Determination of encapsulation efficiency was performed
according to the methodology established by Aizpurua-Olaizola et al. (2016) with modifications. A mass of microcapsules was ground as finely as possible with a mortar. From
this, 3 samples were taken for each of the species and 7.5 mL
of 0.3 M sodium citrate was added to each of them. Then,
the mixtures were subjected to high-frequency stirring with
a digital Ultra Turrax® IKA T18, gradually increasing the
frequency up to 21,000 rpm, in order to obtain a homogeneous mixture. The mixtures were then incubated in an ELMA
Transsonic 820 / H ultrasound bath for 2 h. Once this process
was finished, 15 mL of methanol was added to precipitate
the polymer and obtain an extract of the pigments. Pigment
extracts were evaluated by UV–Vis spectrophotometry in a
Cary 50 conc Varian spectrophotometer (Pittsburgh, USA)
at 400 nm for P. murcianum and 490 nm for T. australis.
The absorbance measurements correspond to the amount of
encapsulated pigment.

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Statistical analysis
Data were evaluated using descriptive statistics using the
Minitab 19 software (Pennsylvania, USA) (mean, standard
deviation (SD), and relative standard deviation (RSD)). Calibrations were established applying a linear regression model.
Comparison of area under curve was done using the GraphPad Prism 6.01 software (San Diego, CA, USA).

Results
Crude extract yields
The culture broths, rich in pigments from both fungi (Fig. 1)
and produced under optimized conditions (Hernández et al.
2019), were treated with Amberlite XAD-16 resin and then
concentrated. The broths yielded 3 mg/mL of yellow and
brown pigments in the case of P. murcianum and 5 mg/mL
of mixture of red and yellow pigments for T. australis.

Separation of fungal extracts by CPC
After subjecting the crude extracts to liquid–liquid extraction with different biphasic solvent systems, the distribution
of pigments between the phases of the same system was
evaluated and the system that showed the best distribution
of pigments was selected. Thus, the system ethyl acetate/
butanol/water was selected for P. murcianum (1:1:2 v/v/v)
and for T. australis (4:1:5 v/v/v). In a second test, a butanol/
water system (1:1 v/v) was used for T. australis, achieving

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a better separation of pigments. Retention of the stationary
phase corresponded to > 80%. After CPC, more than 35 fractions (20 mL) were obtained from the crude extracts of both
fungi, separated in decreasing order of polarity (Figure S1).

Chemical characterization of P. murcianum samples
The chromatographic analysis generated fractions relatively
pure of P. murcianum extracts. Multiple yellow compounds
were detected whose mass–charge ratio values coincided
with the molecular weight of several already known natural
pigments. Most of them correspond to polyketides, such as
azaphilones (Gao et al. 2013). Two samples, selected due
to their purity and yield at CPC-HPLC, were selected for a
product ion scan analysis. In the first sample (P. murcianum
sample 1), the main signal of the chromatogram, at 446 nm
and at retention time of 7.62 min (Fig. 2A), indicated the
presence of compounds that absorb in the visible region.
The spectrum registered in the range of 200–600 nm showed
absorbance peaks mainly in the UV–Vis region. In the near
UV–Vis region, there were peak values at 310, 371, 398, and
446 nm (Fig. 2B), which are characteristics for yellow pigments. Measurements in ESI–MS scan mode reveal values
that could correspond to molecular ions of most pigments in
the sample, where at least three mass values (m/z) of 411.15,
421.15, and 438.10 were observed (Fig. 2C). The MS–MS
spectrum of the ion 421.15 describes at least 6 mass values,
144.30, 212.00, 240.90, 245.00, 255.00, and 257.20, that
could correspond to the fragments formed (Fig. 2D) (Figure S2). In the second sample (P. murcianum sample 2), the
signal of the main pigment obtained from the chromatogram

Fig. 1  Culture broths of the
filamentous fungi P. murcianum
(A) and T. australis (B) grown
in PGY medium, under optimized conditions for production
of extracellular red and yellow
pigments, respectively

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Fig. 2  A UHPLC chromatogram (tR = 7.62 min). B Spectrogram of 200–600 nm. C Mass spectrum and chemical structure of
monascin (numbering detail: 1 = [M-H-CO2 + HCOOH + ­H2O]−,

2 = [M-H + HCOOH + ­H2O]−,
3 = [M-2H + HCOOH + ­2H2O].−).
D MS–MS spectrum from P. murcianum sample 1

at 430 nm shows a maximum at 12.00 min (Fig. 3A). Spectrograms in a range of 200–800 nm showed a peak in the
far UV region. To a lesser extent, there were absorption
peaks in the medium UV region at 278 and 312 nm. Finally,
in the visible region, a notorious maximum of 407 nm,

characteristic for yellow pigments, was observed (Fig. 3B).
UHPLC-MS scan mode measurements showed a spectra
with mass values of 319.10 (Fig. 3C). In the MS–MS spectrum of the main molecular ion, it was 319.10, and at least
14 mass values were obtained 83.30, 93.10, 95.00, 98.00,

Fig. 3  A UHPLC chromatogram (tR = 12.00 min). B UV–VIS spectrogram from 200 to 800 nm. C Mass spectrum and chemical structure of
monashexenone (numbering detail: 1 = [M-H].−). D MS–MS spectrum from P. murcianum sample 2

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106.00, 108.10, 120.70, 123.00, 137.00, 145.20, 149.30,
159.00, 175.00, and 180.90 (Fig. 3D) (Figure S3).

Chemical characterization of T. australis samples
The chromatographic analysis of T. australis extracts showed
the presence of less pure fractions since multiples peaks
with low resolution were observed. This evidenced the
complexity of the chemical composition of the extracts of
this fungal specie. Two pigment fractions were analyzed; in
the first one (T. australis sample 1), a chromatogram with
low resolution among the numerous signals was obtained.
The most notable was at 14.38 min (Fig. 4A). The spectrum showed peak values at 429, 458, and 505 nm (Fig. 4B),
which are characteristics for yellow, orange, and red pigments. UHPLC-MS scan mode measurements indicate a
spectrum with mass values of 548.20 (Fig. 4C). When analyzing the fragments obtained in the MS–MS spectrum, it
was observed that for [M-H]− = 548.20, at least 9 mass values were produced 167.00, 322.10, 338.90, 365.00, 408.90,
463.80, and 479.40 (Fig. 4D) (Figure S4). In the second
sample (T. australis sample 2), the chromatogram showed
a main signal at 9.95 min and 430 nm (Fig. 5A). Likewise,
the UV–Vis spectrograms showed peak in the UV region.

Fig. 4  A Chromatogram obtained by UHPLC (tR = 14.38 min).
B UV–VIS spectrogram from 200 to 800 nm. C Mass spectrum and
chemical structure for atrorosin H (numbering detail: 1 = [M-H]−,

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In the near UV region, there was a low signal at 312 nm
and in the visible region, a peak at 405 nm was observed,
which is characteristic for yellow pigments (Fig. 5B). The
ESI–MS scan mode revealed a mass spectrum with the m/z
value of 281.05 (Fig. 5C). When analyzing the fragments
obtained in the MS–MS spectrum, it was observed that for
the m/z value of 281.05, 14 mass values were produced
138.70, 150.50, 164.70, 170.10, 179.80, 183.00, 185.10,
194.00, 196.90, 207.60, 211.00, 212.90, 236.00, and 237.00
(Fig. 5D) (Figure S5).

Evaluation of antioxidant capacity through ORAC
assay
Antioxidant activity tests were carried out with total extracts
and with selected fractions based on the yield of the separation. In all cases, the antioxidant activity was positive,
obtaining ORAC values of 118.2 μmol TE/g, 2130.40 μmol
TE/g, and 477.50 μmol TE/g for P. murcianum total extracts,
samples 1 and 2, respectively, and 239.2 μmol TE/g,
235.72 μmol TE/g, and 706.36 μmol TE/g for T. australis total extracts, samples 1 and 2, respectively. The major
antioxidant activity over time for fractions is shown in Figure S6. Samples containing the complete fungal pigment

2 = [M + ­H2O]−, 3 = [M + HCOOH + ­H2O].−). D MS–MS spectrum
from T. australis sample 1

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Fig. 5  A Chromatogram obtained by UHPLC (tR = 9.95 min). B UV–
VIS spectrogram from 200 to 800 nm. C Mass and chemical structure spectrum for damnacanthal (numbering detail: 1 = [M-H]−,

2 = [M-H + ­HCOOH]−, 3 = [M-H + M].−). D MS–MS spectrum from
T. australis sample 2

showed a fluorescence decay less abrupt than the Trolox®
standard, with a prolonged effect up to 60 and almost
100 min for P. murcianum and T. australis samples, respectively, and being comparable to the time of the highest concentration of Trolox®.

50%, and 90% of the size distribution and particle size dispersal coefficient (Span) are shown in Table 1, where the
values of d50 for T. australis and P. murcianum were 329.59
and 543.57 µm, respectively.
Results of the encapsulation efficiency for different
batches of fungal microcapsules are shown in Table 2,
attaining an average of 40.3% for T. australis and 24.9% for
P. murcianum.

Microencapsulation of fungal pigment
Several batches containing microcapsules from P. murcianum and T. australis pigments were obtained after the
process of microencapsulation (Fig. 6). The microcapsules
obtained were later characterized through color measurements in CIE L* a* b* coordinates (Table S1).
Using microscopic tools, the shape, diameter, and diameter distribution of the microcapsules obtained from the
selected samples were determined. The shape observed in
all the batches produced was normally irregular cubes and
spheres (Fig. 7). The size of microcapsules obtained was
classified as intermediate, with sizes in the range of 75 and
1000 µm. As per powder characteristics, according to their
fineness, the microcapsules were classified as coarse, d50
sieve opening = 355–1000 µm. Particle diameter at 10%,

Discussion
The CPC technique was appropriate for fungal pigment
separation due to the high retention of the stationary phase,
corresponding to the volume of the organic phase of ethyl
acetate/n-butanol, which remained inside the equipment
upon achieving equilibrium. The total time of this process
was approximately 40 min, being faster than the traditional
processes of column separation which can take several
hours. The great flow rate achieved with the CPC technique
can explain this behavior (Friesen et al. 2015).

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Applied Microbiology and Biotechnology (2022) 106:8021–8034

8029

Fig. 6  Microcapsules powder from T australis (A, B, and C) and P. murcianum (D, E, and F) selected batches

The chemical complexity of the fungal pigments samples was evidenced by the large number of fractions with
similar physico-chemical characteristics obtained, which
was difficult for the chromatographic separation. Our results
strongly suggest the presence of polyketid pigments such
as azaphilones and hydroxyanthraquinoids in the fungal

extracts. These pigments have demonstrated biological activity which represents an interesting potential for different uses
(Frisvad et al. 1990; Visagie et al. 2015).
Measurements in ESI–MS scan mode for P. murcianum
sample 1 revealed mass values that would correspond to
molecular ions of the most abundant pigments of the sample,

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Applied Microbiology and Biotechnology (2022) 106:8021–8034

Fig. 7  SEM micrographs showing microcapsules without pigments (226× magnification) (A); microcapsules of P. murcianum pigments (271×)
(B); microcapsule of T. australis pigments (261×) (C); polymeric network in a microcapsule of P. murcianum pigments (1550×) (D)
Table 1  Particle diameter at 10%, 50%, and 90% of the size distribution and particle size dispersal coefficient (Span) for T. australis and
P. murcianum microcapsules
Sample

Diameter (μm) ± SD
d10

d50

Span
d90

Batch C
178.93 ± 0.15 329.59 ± 0.15 480.25 ± 0.15 0.914
T. australis
Batch E
327.33 ± 0.13 543.57 ± 0.13 759.81 ± 0.13 0.796
P. murcianum

Table 2  Encapsulation efficiency percentage for T. australis and P.
murcianum microcapsules
Species

Batch

Encapsulation
efficiency (%)

T. australis

A
B
C
D
E
F

40.4
39.1
41.5
24.4
28.4
22.0

P. murcianum

where at least three mass values (m/z) of 411.15, 421.15, and
438.10 were observed. It is necessary to consider that these
values may contain adducts with formic acid (HCOOH) and
­H2O due to the high affinity that these pigments would have
with the phases used here. When analyzing the molecular
weights of known pigments, a mass value of 421.15 was
selected. This value is close to the molecular weight of
the yellow pigment monascin (­ C21H26O5, MW = 358.43 g/
mol) found in the species of Monascus and Penicillium
(Mapari et al. 2008). In this case, it would be forming an
adduct with HCOOH and ­H2O, justifying the m/z value of
421 ([M-H + HCOOH + ­H2O]−). The m/z value 411 would
correspond to the compound monascin without the carbonyl group and forming an adduct with HCOOH and two

molecules of ­H2O ([M-H + HCOOH + ­2H2O-CO]−). Likewise, the m/z value 438 could correspond to the molecular
ion of monascin, forming an adduct with HCOOH and two
molecules of H
­ 2O, and with the loss of a second hydrogen
[M-2H + HCOOH + ­2H2O]−. There are some evidences
supporting these observations. For instance, according to
Mapari et al. (2008), the monascin pigment obtained from
the species of Penicillium also forms an adduct with acetonitrile and sodium [M + Na + ­CH3CN]+ = 422.19, showing
absorption peaks at 234, 292, and 394 nm. These data are
similar to that obtained in the present study, although in this
case the adducts would be formed with HCOOH and H
­ 2O.
Another evidence that supports the presence of monascin
is reported by Yang et al. (2018), who describe the formation of a product from monascin with a mass value m/z 331
in ESI ( +) for Monascus species. This is close to the m/z
329 obtained in this analysis. Here, a possible adduct with
HCOOH and H
­ 2O was detected with a m/z 411.15 in the
mass spectrum. The MS–MS spectrum of the ion 421.15
showed at least six mass values that would correspond to the
fragments formed by monascin. All values are theoretically
explained, and the possible structures of the fragments are
proposed in Figure S2.
UHPLC–DAD–ESI–MS scan mode measurements for
P. murcianum sample 2 revealed a spectrum that provide a
mass value of 319.10, corresponding to the main molecular ion in the fraction. Therefore, the molecular weight
should be approx. 320, the value matching with the yellow
monashexenone polyketide ­(C19H28O4, MW = 320.41 g/mol)
found in the species of Monascus (Patakova 2013; Yuliana
et al. 2017). In the MS–MS spectrum of the main molecular ion, it was 319.10, and at least 14 mass values were
obtained. These can be explained through the theoretical
fragmentation of the chemical structure of monashexenone,
described in Figure S3.
UHPLC–MS scan mode measurements for T. australis
sample 1 showed a spectrum with a mass value of 548.20,
which would correspond to the main molecular ion in the
pigment, atrorosin H ­(C29H31N3O8, MW = 549.55 g/mol).

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Applied Microbiology and Biotechnology (2022) 106:8021–8034

This pigment has been described as atrorosins (Isbrandt et al.
2019), a new subgroup of Monascus-type pigments produced
by the species T. atroroseus (Tolborg et al. 2020). These pigments are characterized by having an amino acid integrated
into the pyridine ring that is part of the base ring of nitrogenated azaphilones, which in this case corresponds to histidine (Isbrandt et al. 2019). The value [M-H]− = 548.20 is
close to that described by Tolborg et al. (2020), who obtain
a value of [M + ­H]+ = 550.22 during LC–MS analysis. The
two values could correspond to the molecular weight 549.20,
only differing in the type of ionization used (negative and
positive, respectively). When analyzing the fragments
obtained in the MS–MS spectrum, it was observed that for
[M-H]− = 548.20, at least 9 mass values were produced.
These can be explained by the fragmentation of atrorosin
H, detailed in Figure S4.
The ESI–MS scan mode revealed for T. australis sample
2 a mass spectrum with the m/z value of 281.05, corresponding to the main molecular ion in pigment fraction. This signal may correspond to the natural yellow pigment hydroxyanthraquinoid damnacanthal ­(C16H10O5, MW = 282.25 g/
mol) (Caro et al. 2012). The m/z of 327.05 observed could
be associated to two joined damnacanthal molecules, one
of them without a proton ([M-H + ­M]−). When analyzing
the fragments obtained in the MS–MS spectrum, it was
observed that for the m/z value of 281.05, 14 mass values
were produced, which could correspond to the damnacanthal
molecular fragments described in Figure S5.
The chemical characterization of the main pigments
that are present in extracellular cultures of the filamentous
fungi P. murcianum and T. australis fractioned by CPC is
here described for the first time. In both fungal species, pigments of azaphilone and anthraquinoid class predominated.
In the extracts of P. murcianum, it was possible to identify
compounds responsible for the yellow coloration, such as
monascin and monashexenone. In the case of T. australis, it
was possible to identify the compound responsible for the
red coloration, such as atrorosin H, and compounds responsible for the yellow coloration such as damnacanthal. The
compounds found here have not been previously described
in these fungal species; however, for T. australis, other
azaphilone-type pigments have been reported (Visagie et al.
2015). Additional spectroscopic analysis such as C-NMR
and H-NMR could be performed in the future for definitive
confirmation of proposed compounds.
The analysis of the biological activity of the P. murcianum and T. australis pigments revealed a high antioxidant activity as determined by the ORAC assay. The ORAC
values obtained were comparable with those reported by Jin
and Pyo (2017). In order to relate the chemical nature of the
molecules with the antioxidant effect, several fungal fractions were analyzed in more detail. The Trolox® reagent is
a water-soluble synthetic analog of vitamin E, which acts

8031

exclusively by reducing the phenolic hydroxyl in the chroman ring (Forrest et al. 1994). The fungal pigments analyzed
here could act in similar form, but also act by means of
other mechanisms such as free radical scavenging, being
substrate for radicals and/or chelation of metal ions. The
antioxidant activity in the azaphilones is mainly related with
two different mechanisms: direct reaction between radicals
and hydroxyl groups linked to aromatic rings, and transformation of molecules in pyridines (Chen et al. 2020). The
4-pyridone and its tautomer, the pyridin-4-ol, are characterized as powerful antioxidants, due to the reactivity of vinyl
groups. Although the mechanism is not yet fully clarified,
the structure–activity relationship of pyridines derived
from azaphilones has been demonstrated (Gao et al. 2013;
Ezquerra-Brauer and Chan-Higuera 2021).
The high antioxidant activity (2130 µmol TE/g) detected
in sample 1 of P. murcianum could be related with some of
the mechanisms mentioned above, while being mainly attributed to the presence of monascin pigments. In the case of T.
australis, the antioxidant effect could be mainly related to
anthraquinoid pigments, since sample 2 presented the highest ORAC value (706 μmol TE/g) and contained damnacanthal, which has a proven antioxidant activity (Saha et al.
2013; Li et al. 2017). These results suggest that the great
diversity of molecules present in the fungal extracts may
act in combined ways presenting a synergistic effect in some
cases, or interfering each other in other cases (antagonistic
effect). The latter would explain the notable difference in
ORAC values of purified fractions versus raw extracts of the
fungi. The high antioxidant activity detected in the purified
fungal fractions by ORAC assay suggests the potential of
these natural pigments as innovative functional ingredients
to be used in the cosmetic industry.
The use of microencapsulation allowed the obtention of
homogeneous powders containing fungal pigments, which
could be useful for the cosmetic industry. In the microcapsules, sharp edges and dotted surfaces with small indentations were observed in some cases (Fig. 6). This may be due
to the collapse of the polymeric structure during the lyophilization process. As an automated process, microencapsulation is a fast and simple method to obtain microcapsules of
low size distribution. These characteristics are necessary in
standardized product, since uniform microcapsules ensure a
uniform pigment amount (Swarbrick 2006). The polymeric
materials, sodium alginate, hyaluronic acid, and chitosan,
used in the process are all biodegradable and bio-compatible, allowing also their application as a sustainable product
(Necas et al. 2008; Lupo et al. 2012; Leong et al. 2016;
Shariatinia 2019). Optical microscopy contributed to the
routine monitoring of the experiments carried out to obtain
microencapsulates. On the other hand, scanning microscopy
allowed a more detailed analysis of the shape and structure
of selected samples.

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8032

Applied Microbiology and Biotechnology (2022) 106:8021–8034

Another characteristic to be noted is the already reported
low toxicity of the extracellular pigments produced by T.
australis and P. murcianum. Cytotoxicity tests conducted by
Hernández et al. (2019) provided evidence on the safe nature
of these pigments against mammalian cell lines (HEK293
and NIH/3T3) to which pigments and pigment-leachates
were applied. In this sense, more research including toxicity test oriented to cosmetic uses, such as immunological
test to determine skin hypersensitivity and testing on other
skin cell lines, would be appropriate and highly advisable
for these fungal pigments.
The results obtained in this work contribute to the chemical and biological characterization of new natural ingredients of fungal origin as an alternative to synthetic ingredients and dyes for industrial use, such as in the cosmetics
industry.
Supplementary information The online version contains supplementary material available at https://d​ oi.o​ rg/1​ 0.1​ 007/s​ 00253-0​ 22-1​ 2255-9.
Acknowledgements MAB thanks the support of FONDECYT project
N° 1171857 and FONDEQUIP project N° 130209. VHC thanks the
support of PAI Convocatoria Nacional Subvención a Instalación en la
Academia 2018, 77180054 and Fondecyt 11180030. We appreciate
the valuable contributions of the reviewers of the first version of this
manuscript.
Author contribution PCM: conceptualization, resources, methodology,
software, formal analysis, writing—original, writing—review and editing draft, visualization, project administration. MAL: methodology,
resources, supervision. EPN: methodology, resources, software, validation, formal analysis, visualization, supervision, writing—review and
editing draft. AMH: conceptualization, resources, writing—original
draft, writing—review and editing draft. MAB: methodology, software,
validation, supervision, writing—review and editing draft. VHC: conceptualization, resources, review and editing draft. MFE: methodology,
conceptualization, resources, writing—review and editing draft.
Data availability The datasets generated during and/or analyzed during the current study are available from the corresponding author on
reasonable request.

Declarations
Ethical approval This article does not contain any studies with human
participants or animals performed by any of the authors.
Conflict of interest The authors declare no competing interests.

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Authors and Affiliations
Paulina I. Contreras‑Machuca1,2 · Marcia Avello1 · Edgar Pastene3 · Ángela Machuca4 · Mario Aranda5 ·
Vicente Hernández6,7 · Marcos Fernández2
Marcia Avello
maavello@udec.cl

3

Laboratory of Synthesis and Biotransformation of Natural
Products, Universidad del Bio Bio, Chillán, Chile

Edgar Pastene
edgar.pastene@gmail.com

4

Fungal Biotechnology Laboratory, Department of Plant
Sciences and Technology, Universidad de Concepción,
Campus Los Angeles, Los Angeles, Chile

5

Food and Drug Research Laboratory, Department
of Pharmacy, Faculty of Chemistry and Pharmacy, Pontificia
Universidad Católica de Chile, Santiago, Chile

6

Faculty of Forestry, Universidad de Concepción, Concepción,
Chile

7

Biotechnology Center, Universidad de Concepción,
Concepción, Chile

Mario Aranda
mario.aranda@uc.cl
Vicente Hernández
vhernandezc@udec.cl
Marcos Fernández
marferna@udec.cl
1

Pharmacognosy Laboratory, Department of Pharmacy,
Faculty of Pharmacy, Universidad de Concepción,
Concepción, Chile

2

Pharmaceutical Technology Laboratory, Department
of Pharmacy, Universidad de Concepción, Concepción, Chile

13
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