Screening For Yeast Phytase Leads to the Identification of a New Cell-Bound and Secreted Activity in Cyberlindnera jadinii CJ2

Abstract

Phytic acid is an anti-nutritional compound able to chelate proteins and ions. For this reason, the food industry is looking for a convenient method which allows its degradation. Phytases are a class of enzymes that catalyze the degradation of phytic acid and are used as additives in feed-related industrial processes. Due to their industrial importance, our goal was to identify new activities that exhibit best performances in terms of tolerance to high temperature and acidic pH. As a result of an initial screening on 21 yeast species, we focused our attention on phytases found in Cyberlindnera jadinii, Kluyveromyces marxianus, and Torulaspora delbrueckeii. In particular, C. jadinii showed the highest secreted and cell-bound activity, with optimum of temperature and pH at 50°C and 4.5, respectively. These characteristics suggest that this enzyme could be successfully used for feed as well as for food-related industrial applications.

Full text

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ORIGINAL RESEARCH
published: 24 May 2021
doi: 10.3389/fbioe.2021.662598
Edited by:
Lucia Gardossi,
University of Trieste, Italy
Reviewed by:
Hasan Bugra Coban,
Dokuz Eylül University, Turkey
Farshad Darvishi,
Alzahra University, Iran
*Correspondence:
Concetta Compagno
concetta.compagno@unimi.it
Specialty section:
This article was submitted to
Industrial Biotechnology,
a section of the journal
Frontiers in Bioengineering and
Biotechnology
Received: 01 February 2021
Accepted: 29 March 2021
Published: 24 May 2021
Citation:
Capusoni C, Serra I, Donzella S
and Compagno C (2021) Screening
For Yeast Phytase Leads to the
Identification of a New Cell-Bound
and Secreted Activity in Cyberlindnera
jadinii CJ2.
Front. Bioeng. Biotechnol. 9:662598.
doi: 10.3389/fbioe.2021.662598
Screening For Yeast Phytase Leads
to the Identification of a New
Cell-Bound and Secreted Activity in
Cyberlindnera jadinii CJ2
Claudia Capusoni, Immacolata Serra, Silvia Donzella and Concetta Compagno*
Department of Food, Environmental and Nutritional Sciences, University of Milan, Milan, Italy
Phytic acid is an anti-nutritional compound able to chelate proteins and ions. For
this reason, the food industry is looking for a convenient method which allows its
degradation. Phytases are a class of enzymes that catalyze the degradation of phytic
acid and are used as additives in feed-related industrial processes. Due to their industrial
importance, our goal was to identify new activities that exhibit best performances in
terms of tolerance to high temperature and acidic pH. As a result of an initial screening
on 21 yeast species, we focused our attention on phytases found in Cyberlindnera
jadinii,Kluyveromyces marxianus, and Torulaspora delbrueckeii. In particular, C. jadinii
showed the highest secreted and cell-bound activity, with optimum of temperature and
pH at 50◦C and 4.5, respectively. These characteristics suggest that this enzyme could
be successfully used for feed as well as for food-related industrial applications.
Keywords: phytic acid, yeast, Cyberlindnera jadinii, feed additive, food production, phytase
INTRODUCTION
Phytic acid (myo-inositol 1,2,3,4,5,6-hexakis dihydrogen phosphate) is the main source of stored
phosphorus in grains, oil seeds, and nuts (Mullaney and Ullah, 2003), typically representing up to
60–80% of total phosphorus in seed, and playing an important role during seed germination and
growth (Shi et al., 2005). The presence of phytic acid creates problems in breeding, being feeds
mainly composed by vegetal materials rich in this acid. Polygastric animals are able to degrade
phytate, thanks to their particular gut microbiota (Nakashima et al., 2007), but this process does
not occur in the monogastric ones, like poultry, pigs, fishes, and also humans. Since phytate
cannot be metabolized, feed for monogastric animals are often fortified with inorganic phosphorus,
increasing their final cost. In addition, accumulation of phytic acid has a negative effect on animal
health, because it represents an anti-nutritional and chelating agent, that reduces bioavailability of
proteins and ions like Fe3+, Ca2+, Zn2+, and Mg2+forming insoluble complexes (Reddy et al.,
1982;Coban and Demirci, 2017). The undigested phytate then accumulates in manure and liquid
effluents, leading to phosphorus pollution and water eutrophication. Also for human nutrition,
there is now an increasing attention about these aspects. Phytate degradation in food is mediated
mainly by fermentation processes led by phytate-degrading microorganisms (De Angelis et al.,
2003;Rizzello et al., 2010) or during the food processing by endogenous phytases present in food
Abbreviations: aa, aminoacid; EDTA,ethylene-diamine-tetraacetic acid; OD, optical density; TCA, tricloracetic acid; gDNA,
genomic DNA; U, unit.
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Capusoni et al. Phytase From Cyberlindnera jadinii
matrix (Leenhardt et al., 2005). For the reasons described above,
science community has focused attentions on phytate-degrading
enzymes (Lei et al., 2013;Mrudula Vasudevan et al., 2019;
Pires et al., 2019).
Phytases are a class of enzymes that catalyze the hydrolytic
degradation of phytic acid to free inorganic phosphorus, to yield
lower myo-inositol phosphate esters and, in some case, free
myo-inositol (Vats and Banerjee, 2004). Enzymes described as
phytases show different structures: histidine acid phosphatase
(HAP), βpropeller phytase (BPP), and purple acid phosphatase
(PAP) (Mullaney and Ullah, 2003). The most known and wide
class is HAPs (EC 3.1.3.8). This class is ubiquitous, indeed
HAPs can be found not only in bacteria, yeasts, and filamentous
fungi but also in upper eukaryotes (Lei et al., 2013). All the
proteins belonging to this class maintain two common domains:
a conserved N-terminal heptapeptide active site RHGXRXP (aa
38–44) and a C-terminal catalytically active dipeptide HD (aa
325–326) (Mullaney and Ullah, 2003). Among HAPs exist a
variety of specific activities. Wyss and coworkers analyzed several
fungal phytases dividing them in two different subclasses: one
with broad substrate specificity but low specific activity on phytic
acid (PhyBp), and the second with narrow substrate specificity
but high activity on phytic acid (PhyAp) (Wyss et al., 1999).
Curiously, some organisms as Aspergillus niger possess both
forms (Mullaney and Ullah, 2003).
Although phytases have been reported in a wide range of
bacteria, not many of them have been used so far as feed
supplement, since their neutral/alkaline pH optimum and their
optimal temperature could preclude their activity in these
processes. For industrial applications, the ideal phytase should
display in fact three characteristics: ability to hydrolyze phytic
acid in the upper digestive tract of the animals, resilience up
to 65–90◦C and cheap production cost. In particular, to work
properly in the digestive tract, phytase needs to have a pH
optimum between 3.5 and 5.5 and optimum of temperature in the
range 37–40◦C. Furthermore, phytases should actively work also
at higher temperatures, required in feed production processes,
as during pelletting and heat treatment to control Salmonella
spoilage (Mrudula Vasudevan et al., 2019). In addition, it would
need to be resistant to protease activity and to show low
sensitivity to ions (Wyss et al., 1999).
Addition of phytases is currently not yet applied in human
food production for increasing the mineral bioavailability of the
food. This is mainly due to the fact that, so far, all the commercial
phytase-producing organisms are genetically modified organisms
(GMOs), which are commonly not well accepted for human
food production.
Yeasts are good candidates for phytase production and
some of them have been already characterized (Ragon et al.,
2009;Nuobariene et al., 2011;Greppi et al., 2015;Pires et al.,
2019). These enzymes show different localizations: in some
species, phytases are extracellular enzymes, in others are cell
bound or released in the periplasmic space (In et al., 2009;
Olstorpe et al., 2009;Kaur et al., 2010;Hellström et al., 2015;
Kłosowski et al., 2018).
With the aim to find new phytases with characteristics suitable
for feed and food industrial applications, in this study, we
screened 28 yeast strains isolated from different environments,
terrestrial and marine. We investigated cellular localization
of phytase activity and, for some of them, the effects of
different phosphorus sources on their expression. In addition, we
characterized optimal temperature and pH.
MATERIALS AND METHODS
Yeast Strains
The yeast strains studied in this work (Table 1 and
Supplementary Table 1) belong to CBS collection, UBO
Culture Collection (UBO-CC1), DBVPG collection, and private
collections at the University of Milan (CML and UMY). Yeasts
were stored in YPD-20% (vol/vol) glycerol stocks at −80◦C.
Media and Cultivation
Yeasts were cultivated on different media.
YPD: 10 g/L yeast extract, 20 g/L peptone, 20 g/L glucose.
MMPhy: 20 g/L glucose, 5 g/L (NH4)2SO4, 0.5 g/L
MgSO4∗7H20, 0.11 g/L phytic acid sodium salt hydrate (Sigma
Aldrich, Milano, Italy), trace metals (di-sodic EDTA 15 mg/L,
ZnSO4∗4H2O 4.5 mg/L, MnCl∗4H2O 0.1 mg/L, CoCl2∗6 H2O
0.3 mg/L, CuSO4∗5H2O 0.3 mg/L, Na2MoO4∗2H20 0.4 mg/L,
CaCl2∗2H2O 4.5 mg/L, FeSO4∗7H2O 3 mg/L, H3BO31 mg/L,
KI 0.1 g/L) and vitamins, d-biotin 0.05 mg/L, calcium D-
pantothenate 1 mg/L, nicotinic acid 1 mg/L, myoinositol 25 mg/L,
thiamine hydrochloride 1 mg/L, pyridoxine hydrochloride,
p-aminobenzoic acid 0.2 mg/L) as reported in Merico et al. (2007)
with some modification.
MM-: 20 g/L glucose, 5 g/L (NH4)2SO4, 0.5 g/L MgSO4∗7H2O,
trace metals, and vitamins as in MMPhy.
MMP: 20 g/L glucose, 5 g/L (NH4)2SO4, 0.5 g/L
MgSO4∗7H2O, KH2PO41 g/L, trace metals, and
vitamins as in MMPhy.
MMP/Phy: 20 g/L glucose, 5 g/L (NH4)2SO4, 0.5 g/L
MgSO4∗7H2O, 0.11 g/L phytic acid sodium salt hydrate, KH2PO4
1 g/L, trace metals, and vitamins as in MMPhy.
In all media, pH was adjusted to 4.5 using H2SO4.
Yeast cells were cultivated at 28◦C in a rotary shaker at
150 rpm in bluffed flasks (100 ml) containing 20 ml of medium.
Optical density was monitored at 600 nm (OD600). Cells were
precultured for 24 h in YPD, harvested by centrifugation
at 5,000 rpm, and washed three times with sterile NaCl
solution (9 g/L). Then they were used to inoculate alternatively
MMPhy, MMP, MMP/Phy, and YPD at an initial OD600 = 1
(corresponding to 107cells/ml approximately).
Dry Weight Determination
For dry weight measurements (DW), samples from different
culture conditions were collected (in triplicate at each point).
Cells were filtered through a glass microfiber GF/A filter
(Whatman), washed with three volumes of deionized water and
dried at 100◦C for 24 h.
1http://www.univ-brest.fr/ubocc
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TABLE 1 | Results of phytase screening performed on 28 yeast strains grown on medium containing phytic acid as sole phosphorus source.
Yeast Strain OD600 nm Cell-bound Extracellular
Debaryomyces hansenii MI 1 5.5 BDL BDL
Ciberlindnera jadinii CJ2 28 YES YES
Schizosaccharomyces pombe Y709 4 BDL BDL
Kluyveromyces lactis CBS 2359 3 YES BDL
Kluyveromyces lactis Y1356 1 BDL BDL
Kluyveromyces marxianus Y1058 14 YES BDL
Hanseniaspora uvarum UMY 514 8 YES BDL
Hanseniaspora uvarum UMY 571 14 YES BDL
Saccharomyces cerevisiae CENPK 113 7D 7.78 BDL BDL
Saccharomyces cerevisiae LALVIN T73 8.36 BDL BDL
Brettanomyces bruxellensis CBS 2499 10 YES BDL
Lachancea thermotolerans CBS 6340 8 YES BDL
Torulaspora delbruekii CBS1466 8.56 YES BDL
Candida humilis CBS 5658 8.6 BDL BDL
Candida milleri CBS 6897 11 BDL BDL
Rhodosporidium azoricum DBVPG 4620 12 BDL BDL
Zygosaccharomyces kombutchaensis CBS 8849 7 YES BDL
Kazakistania unispora CML133 3 BDL BDL
Meyerozyma guilliermondii UBOCC-A-214008 26 YES BDL
Meyerozyma guilliermondii UBOCC-A-214143 20 YES BDL
Pichia guilliermondii EX15 UBOCC-A-208004 21 YES BDL
Rhodotolura mucilaginosa UBOCC-A-214025 10 BDL BDL
Rhodotolura mucilaginosa UBOCC-A-214036 7.9 YES BDL
Candida atlantica Mo31 UBOCC-A-208026 25 YES BDL
Candida oceani Mo39 UBOCC-A-208034 4.2 BDL BDL
Debaryomyces hansenii BIO2 UBOCC-A-208002 11 YES BDL
Debaryomyces hansenii Mo40 UBOCC-A-208035 8.2 BDL BDL
Rhodotorula diobovata Mo38 UBOCC-A-208033 12.5 YES BDL
References for strains origin are reported in Supplementary Table 1. BDL, below detection limit.
Phytase Activity Determination
Enzymatic activity was determined on supernatants (for
extracellular activity) and whole cells (for cell-bound activity) of
early exponential phase cultures. The activity was measured by
ortho-phosphate production, following ammonium molibdate
blue method as reported by Schimizu (Shimizu, 1992), with
some modifications.
For extracellular activity determination, cell cultures were
centrifuged at 13,000 rpm and 1 ml of supernatant was added
to 4 ml of buffer composed by 0.2 M Na acetate/acetic acid,
8 mM phytic acid at pH 4.5. To determine cell-bound activity,
a standard amount of cells (corresponding to 10 mg of dry
weight) was collected, washed twice with 0.2 M Na acetate/acetic
acid at pH 4.5, and resuspended in a final volume of 1 ml of
water. Cell suspension was added to 4 ml of buffer in 0.2 M
Na acetate/acetic acid, 8 mM phytic acid at pH 4.5. All buffers
employed to test enzymatic activity were prewarmed at the
reaction temperature. Blank was assembled using 1 ml of water
and 4 ml of 0.2 M Na acetate/acetic acid at pH 4.5, 8 mM phytic
acid, and treated as sample.
For enzymatic activity determination, 5 ml of reaction mixture
were incubated in 15 ml tube at 37◦C and stirred at 300 rpm.
The reaction was immediately stopped (for time 0) and then
stopped after 15, 30, 60, and 120 min by mixing 0.5 ml of
reaction mixture with 0.5 ml of TCA 5% solution. Samples
were centrifuged for 3 min at 13,000 rpm and the supernatants
collected. To determine orthophosphate concentration, 0.4 ml
of supernatant was added to 0.4 ml of molibdate solution.
This solution was prepared daily, by mixing solution A and B
in a ratio of 4:1 (solution A: 2.6% N6H24Mo7O24 ∗4H2O and
5.5% H2SO4; solution B: 4.6% FeSO4∗7H2O). The sample was
incubated 10 min at 25◦C and read against blank at OD700.
Phosphate concentration was determined using a standard curve
for KH2PO4. One unit of phytase is defined as the amount
of protein that hydrolyses 1 µmol of phosphorus/min. Specific
activity is expressed as milliunits per milligram of cell dry weight.
To determine the effect of temperature, samples prepared with
prewarmed buffer (pH 4.5) were incubated at 50 and 60◦C. To
determine the effect of pH on enzymatic activity, pH buffers
were adjusted at pH 4 and pH 5.5, and the reactions incubated
at 37◦C.
Genomic Extraction
To isolate genomic DNA, pellets corresponding to 30 OD of cells
were resuspended in 0.5 ml of 50 mM Tris–HCl, 20 mM M EDTA
at pH 7.5. This suspension was transferred to a precooled tube
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with an equal volume of glass beads (425–600 µm). Mechanical
lysis was performed using a TissueLyser LT (Qiagen) alternating
2 min of agitation at 50 Hz with 1 min in ice for four cycles.
The supernatant was added with 25 µl of SDS 20% (w/v) and
incubated at 65◦C for 30 min. Immediately, 0.2 ml of 5 M
potassium acetate was added and the tubes were placed on ice
for 30 min. Samples were centrifuged at 13,000 rpm for 5 min
and supernatants transferred to a fresh microcentrifuge tube.
The DNA was precipitated by adding 1 vol of isopropanol.
After incubation at room temperature for 5 min, the tubes were
centrifuged for 10 min. The DNA was washed with 70% ethanol
and dissolved in 50 µl of TE RNAse (10 mM Tris–HCl, 1 mM
EDTA, pH 7.5 RNAse 100 µg/ml). Samples were incubated at
37◦C for 30 min.
Strain Identification
gDNA was amplified with Phusion taq polymerase employing
universal primers for amplification on D1/D2 domain of the
26S rDNA (NL1: GCATATCAATAAGCGGAGGAAAAG, NL4:
GGTCCGTGTTTCAAGACGG) 0.2 µM each, 200 µM dNTP,
and MgCl22.5 mM (Fliegerová et al., 2006). PCR amplification
was carried out by denaturing at 98◦C for 7 min, followed
by 30 cycles of denaturing at 98◦C for 30 s, annealing at
52◦C for 30 s, extension at 72◦C for 1 min, and a final
extension at 72◦C for 5 min. The produced PCR amplicon
was sequenced using the Sanger method at Microsynth Seqlab
(Germany), and the strain was identified by the sequence
similarity using basic local alignment search tool against the
NCBI databases2.
Phylogenetic and Bioinformatics
Analysis
Phytase sequence of D. hansenii Mo40 and Bio2 were obtained in
this work. Phytase gene was amplified from gDNA using primers:
Forward: Phy1 CCGACCATGGATGGTATCGATTTCC,
Reverse: Phy2 CATCGGATCCTAATT GTCACCGGA. Primers
were designed based on D. hansenii CBS 767 (GeneID: 2900382;
XP_460696.1). PCR amplification was carried out by denaturing
at 98◦C for 7 min, followed by 30 cycles of denaturing at 98◦C
for 10 s, annealing at 59◦C for 30 s, extension at 72◦C for
45 s, and a final extension at 72◦C for 10 min. PCR amplicons
were sequenced using the Sanger method at Microsynth
Seqlab (Germany).
The aminoacidic sequences (accession numbers on
Supplementary Table 2) were identified through a BLASTp
by XP_460696.1 of D. hansenii CBS 767 against the NCBI
databases. For phylogenetic analysis, multiple alignments
of aminoacidic sequences were performed using MUSCLE
(EMBL-EBI tool on3) and a maximum likelihood tree was
built using Mega X 10.1.74. Analysis of signal secretion
sequence was performed employing SignalIP-5.0 available on
http://www.cbs.dtu.dk/services/SignalP/.
2http://www.ncbi.nlm.nih.gov/BLAST/
3https://www.ebi.ac.uk/Tools/msa/muscle/
4www.megasoftware.net
RESULTS AND DISCUSSION
Growth in Presence of Phytic Acid as
Sole Phosphorus Source
Twenty-one yeast species (28 strains) belonging to Debaryomyces,
Cyberlindnera,Schizosaccharomyces,Kluyveromyces,
Saccharomyces,Brettanomyces,Candida,Torulaspora,
Rhodosporidium,Meyerozyma,Hanseniaspora,Pichia,
Lachancea,Kazakistania, and Rhodotorula genera were
characterized for their ability to grow using phytic acid as
sole phosphorus source (Table 1 and Supplementary Figure 1).
Strain CJ2 was identified in this work as Cyberlindnera jadinii.
All strains were cultivated on medium MMPhy, and their growth
was monitored for 72 h. In parallel, as negative control, growth
on MM- (without any phosphorus source) was carried out,
and no appreciable growth was detected (data not shown). We
avoided to perform screening on solid medium due to ambiguous
results reported sometimes in literature.
All strains except Kluyveromyces lactis Y1356 were able to
grow using phytic acid as sole phosphorus source, but with
variable extent (Table 1). Some strains, like C. jadinii and
Meyerozyma guilliermondii were able to exceed 20 OD after 24 h
of incubation, reaching 26 OD and 25 OD, respectively, after 48 h.
Other strains reached lower OD values after 72 h of incubation,
and K. lactis CBS 2359 duplicated only two times reaching 4 OD
(Table 1 and Supplementary Figure 1). These differences reflect
species-dependent efficiency of phytase activity, as well as species-
specific mechanism of phytic acid hydrolysis. Literature reports
that in some yeasts, like Debaryomyces castellii, phytase is able to
completely hydrolyze phosphate from phytic acid (Ragon et al.,
2008), but others, like in Kodamaea ohmeri are, on the contrary,
not able to perform a complete hydrolysis of this acid, leaving
some phosphate group not bioavailable (Li et al., 2009).
Screening for Phytase Activity
In order to identify enzymes that show characteristics suitable
for feed/food industrial processes, meaning a phytase able to
work in the upper digestive tract of monogastric animals, with
optimal temperature range between 37 and 40◦C and pH range
between 3.5 and 5.5, we decided to screen phytase activity at 37◦C
and pH 4.5. Extracellular and cell-bound activity was assayed on
cells grown using phytate as sole phosphorus source, on MMPhy
medium. To correctly compare phytase activity in various strains,
we expressed the activity as milliunits per milligram d.w., instead
of milliunits per milliliter as often reported in literature. In this
way, we avoided the bias due to different ability of strains to grow
in presence of phytate (Table 1), reaching different amount of
biomass per milliliter.
Under these conditions, we detected extracellular phytase
activity only in C. jadinii (26.25 mU/mgd.w.). On the other hand,
16 out of the 28 tested strains showed a detectable cell-bound
activity (Figure 1), and C. jadinii was identified as the one
with the highest (58.36 mU/mgd.w.). Lower levels of cell-bound
activity were found in fact in the other species (Figure 1), like
Kluyveromyces marxianus (4.17 mU/mgd.w.), Meyerozyma
guilliermondii (10.49 mU/mgd.w.), Pichia guillermondii
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FIGURE 1 | Cell-bound phytase activity (mU/mgd.w.) detected using whole cells grown on medium containing phytate.
(7.1 mU/mgd.w.), Rhodotorula diobovata (7.99 mU/mgd.w.),
and Torulaspora delbrueckii (6.1 mU/mgd.w.).
Analysis of Conserved Domain
Aminoacidic sequences of phytase were recovered from
RefSeq/GenBank database on https://www.ncbi.nlm.nih.gov/,
except for D. hansenii Mo40 and Bio2, which were sequenced
in this work. All strains analyzed possess a protein that shows
a good homology with phytase of D. hansenii Mo40 and
Bio2, and some of them, like M. guilliermondii,C. jadinii,
and K. marxianus, exhibit even two sequences encoding for
this enzyme. In the case of K. unispora, we could not identify
any sequence even if this strain can make a duplication on
MMPhy medium. Probably, this ability is due to the presence of
a phosphatase that does not specifically cleave phosphate from
phytic acid (Table 1).
Generally, HAP phytase brings two common domains: a
conserved N-terminal heptapeptide active site RHGXRXP (aa
38–44) and a C-terminal catalytically active dipeptide HD
(aa 325–326) (Mullaney and Ullah, 2003). In the analyzed
phytase sequences, the C-terminal domain is always conserved.
N-terminal domain is maintained in all except in R. toruloides,
in which the active site shows mismatch XHGHRXP, leading
us to conclude that all belong to HAP family. These sequences
were used to build a phylogenetic tree (Figure 2). In the red
box, we included sequences that contain a signal peptide for
protein secretion. Literature data (Li et al., 2009) report that
K. ohmeri enzyme contains this sequence, but the tool we
used was not able to recognize it. As reported in Figure 2, all
sequences showing a signal peptide can be clustered, suggesting
that extracellular localization can be phylogenetically related. The
fact that we detected extracellular activity only in C. jadinii,
could probably depend on the use of culture supernatants
without any step to concentrate it. It is possible to speculate
that extracellular phytase activity in other strains was too low
to be detected. This hypothesis is corroborated also by studies
that report in other species extracellular activity detected only in
concentrated samples (Olstorpe et al., 2009;Ragon et al., 2009;
Hellström et al., 2015).
Regulation of Phytase Expression
To carry out this investigation, we selected three species,
C. jadinii,K. marxianus, and T. delbrueckii that showed
high phytase activity and could be interesting for food-related
applications. C. jadinii and K. marxianus are in fact included in
QPS EFSA list (Koutsoumanis et al., 2019), and T. delbrueckii is a
wine starter with commercial name (BIODIVATM—Lallemand).
Phytase activity was analyzed by cultivating yeast cells in the
presence of different phosphorus sources: phytic acid (in MMPhy
medium), phosphate salt (KH2PO4in MMP medium), mixture
of phytic acid and phosphate salt (in MMP/Phy medium), and
also in rich medium (YPD). This allowed to understand the
regulation of phytase expression based on the type of phosphate
available (Table 2).
When C. jadinii cells were cultivated in medium containing
phytic acid as the sole phosphorus source, we detected
58.36 mU/mgd.w.of cell-bound activity (Table 2) and
26.25 mU/mgd.w.of extracellular activity. These activities
were not appreciable when inorganic phosphate was the only
source (MMP), suggesting that under this condition, the enzyme
was not expressed. In addition, the concomitant presence of
phytic acid and KH2PO4(MMP/Phy medium) was not able
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FIGURE 2 | Phylogenetic tree of phytase aminoacidic sequences. The red rectangle includes proteins that contain a signal sequence for secretion (detected using
SignalIP-5.0).
to induce phytase activity (Table 2). On the contrary, when
C. jadinii was cultivated in the presence of organic phosphate
(YPD medium), a reduced level of cell-bound activity was
detected: it decreased in fact from 58.36 mU/mgd.w., measured
TABLE 2 | Effects of phosphorus source on phytase activity.
YPD MMPhy MMP MMP/Phy
C. jadinii 2.02 ±0.4 58.36 ±6.24 BDL BDL
T. delbrueckii 2.39 ±0.29 6.1 ±1.25 BDL BDL
K. marxianus 1.71 ±0.35 4.17 ±1.17 BDL BDL
Cell-bound activity (mU/mgd.w.) was assayed at 37◦C, pH 4.5. BDL, below
detection limit.
in MMPhy, to 2.02 mU/mgd.w.in YPD (Table 2). Under this
condition, extracellular activity was under the detection limit.
The same behavior was observed in T. delbrueckii and
K. marxianus. The highest activities (6.1 and 4.17 mU/mgd.w.,
respectively) were detected in the presence of sole phytic acid
and decreased in YPD (Table 2). As observed in C. jadinii, also
in T. delbrueckii and in K. marxianus, the presence of phosphate
inhibited expression of phytase, being no activity detectable in
cells growing in MMP as well as in MMP/Phy.
In conclusion, a high level of phytase activity can be
expressed only when cells grow using phytate as sole phosphorus
source. On the contrary, the presence of inorganic phosphate
completely inhibits expression of phytase. This happens in
fact also in medium with concomitant presence of phosphate
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Capusoni et al. Phytase From Cyberlindnera jadinii
FIGURE 3 | Effects of temperature on phytase activity, measured at pH 4.5. (A) Cell-bound (full line) and extracellular (dotted line) activity detected on C. jadinii.
(B) Cell-bound activity detected on T. delbruekii and K. marxianus.
and phytic acid. This indicates that sole presence of phytic
acid is not enough to induce phytase activity, and lead us to
conclude that lack of inorganic phosphate is requested for its
expression. The presence of low activity in cells grown in medium
not containing phosphate salt, namely YPD, corroborates
this hypothesis.
An analogous phenomenon has been observed previously
by Olstorpe and colleagues in 2009 (Olstorpe et al., 2009).
They reported that in some Candida species, in Pichia anomala
and in S. cerevisiae, phytase activity was repressed in presence
of inorganic phosphate, but this did not occur in other
species like Arxula adeninivorans, as well as in Cryptococcus
laurentii (Pavlova et al., 2008). By employing mutagenesis, an
improved strain with reduced phosphate repression was obtained
in Pichia kudriavzevii (Qvirist et al., 2017). Understanding
which role plays phosphorus source on expression of phytase
activity is pivotal for set-up of industrial processes, due
to the fact that feed/food matrices as well as cultivation
media based on agrifood wastes can contain different types
of this nutrient.
Characterization of Phytase Activity
Phytases suitable for feed/food-related processes need to work
under conditions present in the upper digestive tract of
monogastric animals, and/or need to be resilient during
production processes. To work in the digestive tract, a
good phytase should exhibit a pH optimum between 3.5
and 5.5 and high activity at 37◦C. Resilience at higher
temperature can be requested because heat treatments are
commonly adopted to contain spoilage and during pelleting
processes. This treatment permits also the incorporation of
ingredients in the feed to “lock” the feed mixture. Unfortunately,
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Capusoni et al. Phytase From Cyberlindnera jadinii
FIGURE 4 | Effect of pH on phytase activity detected in C. jadinii. Activity was
measured at 37◦C.
heat treatment could reduce phytase activity, and for this
reason, it is important to select a thermostable enzyme
(Mrudula Vasudevan et al., 2019).
In order to select enzymes suitable for these applications, the
effects of temperature and pH were investigated (Figures 3,4).
The optimal temperature for phytase activity in C. jadinii
was found to be 50◦C. When the assay was performed at
this temperature, cell-bound activity reached 146 mU/mgd.w.
and extracellular activity was 51.95 mU/mgd.w.At 60◦C, the
values for cell-bound and extracellular activity were lower:
105.2 and 37.2 mU/mgd.w., respectively (Figure 3A). Similar
results were obtained for K. marxianus. Also in this case, the
optimum of temperature was observed at 50◦C, with activity
of 7.11 mU/mgd.w.A similar temperature activity profile was
found for phytase from Rhodotorula mucilaginosa JMUY14 (Yu
et al., 2015). On the other hand, in T. delbrueckii, the highest
activity of 6.1 mU/mgd.w.was detected at 37◦C (Figure 3B).
Data reported in Figure 3 lead us to hypothesize that phytase
activity in C. jadinii could be suitable for industrial purpose. High
phytase activity detected even at 60◦C suggests that this enzyme
could be resilient also at high temperatures used during feeds
production. In addition, at 37◦CC. jadinii phytase activity is
higher in comparison with K. marxianus and T. delbrueckii ones.
With the aim to investigate the effect of pH on C. jadinii
activity, we selected temperature of 37◦C (Figure 4). Even if
this temperature is not the optimum for C. jadinii, it is the
one requested to work at gastric level. As reported in Figure 4,
pH optimum for this phytase was 4.5, similarly to phytases
found in other yeasts (In et al., 2009;Caputo et al., 2015;
Ogunremi et al., 2020). As reported by Lei et al. (2013), this is
an important characteristic for the development of enzymes as
feed/food additive.
Comparing our results with some data reported in literature,
it is possible to observe that phytase activity found in C. jadinii
could be promising for future applications (Supplementary
Table 3). To the best of our knowledge, C. jadinii cell-bound
activity is one of the highest observed on cells grown in mineral
media with phytate as sole phosphorus sources.
In conclusion, we think that these results found for C. jadinii
phytase activity could represent a good starting point to
set-up optimization of cultural conditions, in order to improve
phytase production. As reported, with a statistical approach
for medium optimization, phytase production can be easily
increased (Puppala et al., 2018). The productivity of phytase
using Aspergillus niger under submerged fermentation conditions
was improved by 3.97 times employing a statistical media
optimization strategy and Box-Behnken experimental designs
(Shah et al., 2017). Optimization of temperature, pH, and
aeration allowed the successful production of phytase with
A. ficuum in submerged fermentation as opposed to the
traditional solid-state fermentation (Coban and Demirci, 2014).
In S. cerevisiae modulation of media components, like addition
of magnesium sulfate, manganese sulfate, and ferrous sulfate, and
scaling up in 10 L fermenter could increase phytase activity from
45 to 164 mU/mgd.w..InK. marxianus, phytase activity could be
easily increased adding to fermentation media cheap substrates
rich in phytic acid like rice bran (Pires et al., 2019). Similar
behavior was observed in W. anomalus, where the presence of
cane molasses in media can increase enzymatic activity from 6
up to 176 mU/mgd.w.reducing enzyme production cost from
0.25 to 0.006 £/1,000 U (Vohra and Satyanarayana, 2004). In
our case, performing media optimization could be the right
approach in order to increase phytase productivity, reducing its
production cost.
Furthermore, Cruz and colleagues demonstrated that biomass
of C. jadinii can partially replace feed protein content (generally
consistent in soybean meal, fish meal, rapeseed meal) in swine
and poultry formulation (Cruz et al., 2020a,b). The possibility
to have active enzymes in feed could be very appealing in order
to decrease their phytate content. This phenomenon has been
observed in W. anomalus, whose biomass-containing phytase
was added to aquaculture feed (1,000 U/kg feed), with results
comparable with commercial phytase (Vohra et al., 2011). The
safety statement of C. jadinii could open the possibility for
applications of this no-GMO microorganism also in food-related
processes to produce functional foods (Handa et al., 2020).
CONCLUSION
The combined effect of phytate as antinutritional factor and as
cause of environmental pollution makes phytase an industrially
interesting target. We identified new phytase activities in “safe”
yeasts, like C. jadinii,K. marxianus, and T. delbrueckii. In
particular, C. jadinii shows the highest enzymatic activity
localized both extracellularly and cell-bound. Our results suggest
that this phytase is suitable as additive in feed/food-related
processes. Indeed, its activity showed characteristics in terms of
temperature and pH suitable to work efficiently under conditions
compatible with the upper digestive tract of monogastric animals
as well as to be used in feed industrial production processes. The
safety statement of C. jadinii could open the possibility for its
application to reduce phytate content in food matrices.
Furthermore, a released phytase significantly increases the
interaction between phytate present in food matrix and the
enzyme. In addition, an extracellular enzyme improves greatly
the downstream during industrial production.
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Capusoni et al. Phytase From Cyberlindnera jadinii
DATA AVAILABILITY STATEMENT
The original contributions presented in the study are included
in the article/Supplementary Material, further inquiries can be
directed to the corresponding author/s.
AUTHOR CONTRIBUTIONS
ClC: investigation and writing the original draft. IS: investigation
and review and editing the draft. SD: investigation. CoC:
conceptualization, supervision, and review and editing the
draft. All authors contributed to the article and approved the
submitted version.
SUPPLEMENTARY MATERIAL
The Supplementary Material for this article can be found
online at: https://www.frontiersin.org/articles/10.3389/fbioe.
2021.662598/full#supplementary-material
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Conflict of Interest: The authors declare that the research was conducted in the
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potential conflict of interest.
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