Available via license: CC BY 4.0
Content may be subject to copyright.
Int. J. Mol. Sci. 2022, 23, 15253. https://doi.org/10.3390/ijms232315253 www.mdpi.com/journal/ijms
Article
Biodegradation of Pine Processionary Caterpillar Silk Is
Mediated by Elastase- and Subtilisin-like Proteases
Alba Diez-Galán, Rebeca Cobos *, Ana Ibañez, Carla Calvo-Peña and Juan José R. Coque *
Instituto de Investigación de la Viña y el Vino, Escuela de Ingeniería Agraria, Universidad de León,
24009 León, Spain
* Correspondence: rebeca.cobos@unileon.es (R.C.); jjrubc@unileon.es (J.J.R.C.)
Abstract: Pine processionary caterpillar nests are made from raw silk. Fibroin protein is the main
component of silk which, in the case of pine processionary caterpillar, has some unusual properties
such as a higher resistance to chemical hydrolysis. Isolation of microorganisms naturally present in
silk nests led to identification of Bacillus licheniformis and Pseudomonas aeruginosa strains that in a
defined minimal medium were able to carry out extensive silk biodegradation. A LasB elastase-like
protein from P. aeruginosa was shown to be involved in silk biodegradation. A recombinant form of
this protein expressed in Escherichia coli and purified by affinity chromatography was able to effi-
ciently degrade silk in an in vitro assay. However, silk biodegradation by B. licheniformis strain was
mediated by a SubC subtilisin-like protease. Homologous expression of a subtilisin Carlsberg en-
coding gene (subC) allowed faster degradation compared to the biodegradation kinetics of a
wildtype B. licheniformis strain. This work led to the identification of new enzymes involved in bio-
degradation of silk materials, a finding which could lead to possible applications for controlling this
pest and perhaps have importance from sanitary and biotechnological points of view.
Keywords: silk; pine processionary; Bacillus; Pseudomonas; biodegradation; proteases; elastase;
subtilisin
1. Introduction
Silk is one of the most abundant naturally derived polymers. In fact, silk fibers are a
common material naturally produced by a wide variety of arthropods that use them to
build their cocoons (silkworms), webs (spiders) and nests (honeybees, wasps, and butter-
flies, among others) [1]. Silk production by the Bombyx mori silkworm has developed into
a major textile industry producing around 120,000 metric tons of silk per year, with the
primary manufacturers located in China, India and Japan [1]. We know that most of the
113,000 known species of Lepidoptera can produce silk [2]. One species of silk-producing
lepidopteran is the pine processionary moth (Thaumetopoea pityocampa). This nocturnal
lepidopteran is of particular interest because it has become one of the most harmful pests
affecting forests of several Pinus, Cedrus and Pseudotsuga species in many countries
around the Mediterranean basin. Pine processionary threatens the ecology and sustaina-
bility of pine and cedar forests in many countries in central-southern Europe and North
Africa [3,4]. The problem is worsening because in recent years, and as a putative conse-
quence of global warming, a latitudinal and altitudinal expansion of the pine procession-
ary has been reported in Europe. As a consequence, this pest is moving to the north of
Europe and it has already been detected in Switzerland and Germany [5], threatening
their forests in geographical areas hitherto free of this pest. Furthermore, this species is of
interest because of the severe allergic reactions it can cause in humans, domestic pets and
other animals, even resulting in death, which increases the interest in its control [6].
Citation: Diez-Galán, A.; Cobos, R.;
Ibañez, A.; Calvo-Peña, C.; Coque,
J.J.R. Biodegradation of Pine
Processionary Caterpillar Silk is
Mediated by Elastase- and
Subtilisin-like Proteases. Int. J. Mol.
Sci. 2022, 23, 15253. https://doi.org/
10.3390/ijms232315253
Academic Editor: Hitoshi Sashiwa
Received: 4 November 2022
Accepted: 1 December 2022
Published: 3 December 2022
Publisher’s Note: MDPI stays neu-
tral with regard to jurisdictional
claims in published maps and institu-
tional affiliations.
Copyright: © 2022 by the authors. Li-
censee MDPI, Basel, Switzerland.
This article is an open access article
distributed under the terms and con-
ditions of the Creative Commons At-
tribution (CC BY) license (https://cre-
ativecommons.org/licenses/by/4.0/).
Int. J. Mol. Sci. 2022, 23, 15253 2 of 23
In winter the pine processionary larvae feed voraciously on coniferous needles, caus-
ing severe defoliation which results in both a decrease of the tree’s growth rate and a de-
crease in its annual diameter increment.
Furthermore, defoliated trees can become highly prone to attack by other insects and
pests, resulting in a higher mortality rate in the affected forests [7]. Typically, in winter
the larvae build and live inside silk nests which they leave during the night for feeding on
pine foliage. During the day the larvae rest within these large communal “nests” con-
structed of silken filaments. These nests are firmly attached to the branches of the host tree
and accommodate 50 or more caterpillars and contain a lot of debris in the form of broken
pine needles, wood fragments, fecal matter, and body hairs shed by the caterpillars. The
nests usually have thick, resistant silk walls, providing an additional protection to the
larvae against weather conditions, potential predators and chemical treatments.
Natural silk is characterized by high strength, elasticity, thermal insulation and hy-
groscopicity (in natural conditions silk contains approximately 11% moisture, and it can
have a moisture content of 30% without feeling wet). Silkworm raw silk consists of protein
nanofibers–fibroins–stuck together in groups of 2. Sericin protein envelops the nanofibers
and acts as a glue that holds them together [8,9]. Fibroin makes up 70–75% of raw silk
whereas sericin represents around 25–30%. Sericin is a protein composed of 18 different
amino acids of which serine is the most abundant, representing around 32% of the total
amino acids. Silkworm fibroin is composed of two protein chains, heavy-chain (H-fibroin)
with a molecular weight of approximately 350 kDa and light chain (L-fibroin, Mw~26 kDa)
covalently linked by a disulfide bond at the carboxy-terminus of the two subunits. Disul-
phide bridges are responsible for fiber strength. There are also hydrogen bonds within
and between the molecules. The complete amino acid sequence of the B. mori fibroin heavy
chain is composed of a highly repetitive (Gly-Ala)n sequence motif and tyrosine-rich do-
mains [8,9]. In fact, around 90% of it consists of four amino acids: alanine, glycine, serine,
and tyrosine [10].
Basic characterization of silk fibroin from pine processionary was carried out a long
time ago by Shaw and Smith [11], indicating some typical traits as compared to other silk
fibroins, such as a higher content of amino acids with hydrophilic side-chains, or a higher
resistance to acid hydrolysis. Unfortunately, no further studies on this type of silk have
been carried out to date.
Microbial degradation of other types of silk, particularly silkworm silk (B. mori), is
well documented. Strains of Burkholderia cepacia [12] and Variovorax paradoxus [13] are able
to biodegrade silkworm silk. Microorganisms probably assimilate sericin more easily than
fibroin and it is known that decomposition of sericin primarily involves proteolytic en-
zymes [13]. Two unidentified species of bacteria (belonging to the Streptomyces genus)
have also been reported to produce proteases hydrolyzing a variety of insoluble proteins,
including silk [14,15]. In vitro tests have also confirmed the degradation of fibroin by pro-
tease XIV from Streptomyces griseus [16] and protease XXI from Streptomyces sp. [17]. Both
proteases are actually a mix of proteolytic enzymes including at least three different pro-
teases, of which at least one is an extracellular serine protease. Unfortunately, no infor-
mation is available in the literature indicating the microbial degradation of pine proces-
sionary silk.
Current control strategies employed to suppress the populations of this detrimental
pine tree pest are primarily based on treatments with chemical pesticides and biopesti-
cides, natural compounds and predators, or biocontrol agents, and also the removal of
egg packages, enhancement of natural enemies’ activities, and destruction of winter nests
made of silk [4,18]. In this context, any putative technology that contributes to the degra-
dation and/or disorganization of the nests could be an effective aid in the control of this
plague. This work addresses the isolation and characterization of bacteria with the ability
to degrade raw pine processionary silk and identification of some proteases involved in
the biodegradation process.
Int. J. Mol. Sci. 2022, 23, 15253 3 of 23
2. Results
2.1. Structure of Pine Processionary Raw Silk
Electron microscopy analysis confirmed that the basic structure of raw pine proces-
sionary silk consists of 2 long fibroin nanofibers running in parallel (Figure 1A) which are
associated to form a more complex network of parallel fibers. Both fibroin fibers are held
together by a sericin layer that acts as a glue (Figure 1B,C), with a structure very similar
to that described for silkworm silk. The cross-section of the nanofibers is ovoid and their
average dimensions are 8–10 μm × 4–5 μm (Figure 1B,C).
Figure 1. Scanning electron microscopy (SEM) visualization of fibers of raw pine processionary silk
(X 2000) (A). The basic unit is formed by 2 fibroin nanofibers arranged in parallel and laterally as-
sociated with other fibers of the same nature (A); cross sections of silk basic units observed by trans-
mission electron microscopy (TEM) at 6000 magnification (B) and 10,000 magnification (C). Note
the sericin layer (s) acting as a glue that holds the fibroin nanofibers (f) together.
2.2. Amino Acid Analysis of Silk
Acid hydrolysis of raw silk allowed the detection of 9 amino acids, listed in decreas-
ing concentration: glycine (14.85 ± 0.42 mM), serine (9.19 ± 0.10 mM), alanine (5.72 ± 0.33
mM), aspartic acid (5.68 mM ± 0.24 mM), glutamic acid (2.14 ± 0.14 mM), tyrosine (1.64 ±
0.08 mM), arginine (1.37 ± 0.03 mM), leucine (1.32 ± 0.11 mM) and valine (0.87 ± 0.05 mM).
Some traces of cysteine and phenylalanine/isoleucine mixture (both amino acids coeluted
with the chromatographic conditions used) were also detected although they could not be
quantified.
2.3. Isolation of Silk-Degrading Bacterial Strains
In the initial experiment carried out in a poorly defined minimal medium to force silk
biodegradation, no obvious signs of degradation were visually observed during the first
60 days of incubation. Around day 75 some evident degradation and disorganization of
the silk ball began to be observed, which became more evident around day 90 of incuba-
tion, when only a few loose silk threads remained in suspension. In order to enrich the
sample in putative silk degrading microorganisms, a new batch of silk was reinoculated
with 0.5 mL of the initial culture under the same conditions as the initial isolation but
including glucose at a final concentration of 0.1%. This enrichment step was repeated
twice. In this case we observed a clear acceleration of the degradation process which was
completed on average 10–12 days after inoculation of the sample. Plating serial dilutions
of the enrichment cultures on PCA allowed us to isolate a total of 5 different bacterial
strains based on easily observable microscopic and macroscopic traits of the colonies.
A new sterile silk degradation experiment was performed for each of the isolated
bacteria in pure culture, and we observed that only 2 out of the 5 isolated bacteria showed
Int. J. Mol. Sci. 2022, 23, 15253 4 of 23
clear silk degradation capability. Partial sequencing of their 16S rDNA allowed us to iden-
tify these isolates as Pseudomonas aeruginosa IIVV-SD1 and Bacillus licheniformis IIVV-SD3
(Supplementary material. Figure S1). Strains were named as IIVV by “Instituto de Inves-
tigación de la Viña y el Vino”, the research center in which they were isolated and named
as SD referring to their “Silk Degrading” capability.
Both strains were able to achieve extensive biodegradation of silk in minimal me-
dium, which indicated that silk could be used as both a carbon and nitrogen source. The
observation of liquid cultures under optical microscopy revealed that both isolates with
degradation capability were able to intensively colonize and attach to the surface of the
silk fibers (Figure 2A), producing an evident disorganization and degradation of the fi-
bers. At some points fibers experienced a considerable decrease in thickness, which could
even cause them to break into shorter fragments.
Figure 2. Observation of a P. aeruginosa pure culture under optical microscope (40X) in the presence
of raw silk. (A) 12 h of incubation; (B) 48 h of incubation. Note the intense bacterial colonization
observed on the surface of silk fibers in the 48-hour culture, where silk biodegradation was already
apparent. At some points fibers experienced a considerable decrease in thickness (points marked
with arrows), which could cause the fibers to break into shorter fragments.
Silk biodegradation was confirmed by SEM visualization (Figure 3). Silk biodegrada-
tion seemed to start with removal of the outer sericin layer (Figure 3C,D). After that, bac-
terial cells could gain access to the inner fibroin structure to achieve extensive biodegra-
dation with concomitant loss of fiber structure (Figure 3E,F).
Int. J. Mol. Sci. 2022, 23, 15253 5 of 23
Figure 3. Scanning electron microscopy (SEM) visualization of raw pine processionary silk filaments
and their successive degradation over time mediated by a liquid culture of B. licheniformis. Silk na-
tive structure at 0 h of incubation (X2000) (A); partial removal of the sericin surface layer observed
at 24 h of incubation (X2000) (B); almost complete elimination of the sericin layer detected at 36 h of
incubation and presence of some bacterial cells (b, indicated by arrows) attached to the underlying
fibroin fibers (X5000) (C) (X3000) (D); extensive degradation with loss of fibroin fiber structure ob-
served after 48 h of incubation (X5000, (E)) (X2000, (F)).
2.4. Analysis of Silk Biodegradaton by B. licheniformis and P. aeruginosa in Liquid Media
The ability of both B. licheniformis IIVV-SD3 and P. aeruginosa IIVV-SD1 strains to
biodegrade silk in liquid media was tested in in vitro experiments run for 30 days (Figure
4) in a minimal medium containing sterile silk. Under these experimental conditions we
observed the release of amino acids into the culture supernatant, indicating that both bac-
terial strains were able to biodegrade silk. B. licheniformis growth proceeded quickly over
the first 2 days of the experiment reaching a maximum growth around day 14 with an
average of 1.13 × 108 cfu/mL (Figure 4A). The amino acid release by B. licheniformis in-
creased rapidly during the first 6 days of the experiment, later increasing more slowly
until around day 22. From that day on, the concentration of amino acids in the culture
supernatant tended to remain constant (Figure 4B).
Int. J. Mol. Sci. 2022, 23, 15253 6 of 23
Figure 4. Growth rate of P. aeruginosa IIVV-SD1 and B. licheniformis IIVV-SD3 strains in a minimal
medium supplemented with raw silk (A), and biodegradation of pine processionary raw silk esti-
mated by release of L-Leucine equivalent (mM) into the culture medium (B). Bacterial cultures were
run in parallel to a negative control of raw silk incubated in a non-inoculated minimal medium. The
results shown are the average of two different experiments performed in triplicate.
The behavior of P. aeruginosa showed some remarkable differences. Growth under
the experimental conditions used was slower and more sustained during the first 12 days
of the experiment, reaching maximum growth around day 16, with a mean value of 1.58
× 109 cfu/mL, an order of magnitude higher than that detected for B. licheniformis (Figure
4A). However, despite faster P. aeruginosa growth, silk degradation, and therefore the re-
lease of amino acids into the culture supernatant, was slower. It increased slowly and
steadily during the first 10 days of incubation, increasing more rapidly between days 10
and 20. Beyond day 20, the concentration of amino acids in the supernatant tended to
stabilize (Figure 4B).
2.5. Genome Analysis of B. licheniformis IIVV-SD3: In Silico Analysis to Identify Putative
Proteases Involved in Silk Biodegradation
The draft genome sequence of B. licheniformis IIVV-SD3 presented an estimated ge-
nome size of 4,391,155 bp representing a coverage of 92.65% of the reference genome. It
contained 5440 protein-coding genes, of which 4222 were annotated (77.61%). Of the an-
notated genes, functional annotation of the GO terms was found for 3647 (86.38%) and of
InterPro terms for 3973 (94.1%) genes.
In an attempt to identify putative enzymes involved in silk degradation we focused
our attention on the detection of genes encoding proteases/peptidases with an extracellu-
lar or cell wall location. Prediction of their putative cellular localization was very interest-
ing since we could assume that silk-degrading proteases would be mostly extracellular or
associated to cell wall.
Thus, a total of 6 peptidases had a possible cell wall localization whereas 20 pepti-
dases were classified as extracellular (Supplementary material. Table S1). Of these, a total
of 6 appear to be peptidases associated with wall peptidoglycan processing; most were
Ser-type D-Ala-D-Ala-carboxypeptidases (proteins number 5, 6 and 20 in Table S1), and
Cys peptidoglycan endopeptidases (proteins 11, 12 and 15. Table S1) and therefore they
were initially discarded as possible candidates involved in silk degradation. Other candi-
dates were discarded according to their biological activity (Glutathione hydrolase proen-
zyme; protein number 1), or their minor character (Proteins 4 and 14—Minor extracellular
Int. J. Mol. Sci. 2022, 23, 15253 7 of 23
proteases Epr; protein 9—minor extracellular protease vpr). Among the remaining extra-
cellular proteins, protein 13 (subtilisin Carlsberg Ser-endopeptidase) particularly at-
tracted our attention because it is an extracellular alkaline serine protease, catalyzing the
hydrolysis of proteins and peptide amides that shows high specificity for aromatic and
hydrophobic amino acids at the P1 position of the substrate [19,20]. According to Shaw
and Smith [11], and our own results, aromatic and hydrophobic amino acids are abundant
in pine processionary fibroin and therefore this protein may contain a high number of
potential cleavage sites that would allow for its efficient degradation. Subtilisin Carlsberg
also exhibits a number of other attractive properties, such as high thermostability, wide
range of pH compatibility, and broad specificity [20], that make it an ideal candidate for
its putative involvement in silk degradation, so we decided to focus on it for further stud-
ies.
A single copy of a gene encoding a subtilisin Carlsberg was detected in the genome
of B. licheniformis IIVV-SD3 strain (see Supplementary material Table S2 for the nucleotide
sequence of the subC gene and the corresponding amino acid sequence of SubC protein).
Homologous genes are called subC (from subtilisin Carlsberg), apr_2 or apr (from alkaline
protease), or aprE (from alkaline protease E or subtilisin E) by different authors. We de-
cided to adopt the terminology suggested by the Uniprot database (https://www.uni-
prot.org/uniprotkb/P00780/entry accessed on 24 October 2022), accepting the designation
subC (and using apr as its synonym). Thus the B. licheniformis IIVV-SD3 strain subC gene
encoded a protein of 379 amino acids with a molecular weight of 38,860 Da and an 8.73
isoelectric point. The corresponding SubC protein exhibits a 100% amino acid identity
with other B. licheniformis proteins annotated as keratinase (AFT92040.1) and S8 family
peptidase (WP_025808265.1), and slightly lesser identities (99.47% to 99.74%) with several
other proteins annotated as subtilisin Carlsberg (P00780.2, QNT35445.1), protease
(ABU68339.1), keratinase (AAS86761.1), or S8 family peptidase (WP_186442051.1).
2.6. Genome Analysis of P. aeruginosa IIVV-SD1: In Silico Analysis to Identify Putative
Proteases Involved in Silk Biodegradation
The draft genome sequence of P. aeruginosa IIVV-SD1 presented an estimated genome
size of 6,378,951 bp representing a coverage of 92.27% of the reference genome. It con-
tained 5988 protein-coding genes, of which 4463 were annotated (74.53). Of the annotated
genes, functional annotation of the GO terms was found for 4192 (93.93%) and of InterPro
for 4364 (97.78%) genes.
To identify putative enzymes involved in silk degradation, again we were interested
in predicting their putative cellular localization since we assumed that the proteases re-
sponsible for silk degradation should be mostly extracellular or cell wall-associated. A
total of 25 peptidases had a possible cell wall localization, perhaps located at the level of
the outer cell membrane and/or in the periplasmic space. A total of 9 peptidases were
classified as extracellular (Supplementary material. Table S3). Among them two proteins
caught our attention, namely the metalloendopeptidases LasA (protein 6 in Table S3) and
LasB (protein 2 in Table S3). The latter, also known as elastase or pseudolysin, was partic-
ularly interesting since it is a widely studied protease with broad specificity although it
favors hydrophobic or aromatic amino acid residues, with preference to Phe and Leu at
the P1′ position. LasB also cleaves proteins with Gly at the P1 position, such that the pre-
ferred cleavage sequence is P1Gly-P1′(Leu/Phe)-P2′Ala [21]. Interestingly, porcine elastase
exhibits a different cleavage pattern, mainly cutting after Ala, Gly, Leu, Ile, Ser and Val
residues [22]. As we have previously indicated, according to Shaw and Smith [11] and our
own results, Gly, Ser and Ala are highly abundant, with Leu and Phe present in a lower
amount, in raw pine processionary silk and therefore silk proteins may contain a high
number of potential cleavage sites that allow for efficient degradation by both P. aeru-
ginosa and porcine elastases. P. aeruginosa LasB elastase is the predominant protease in the
P. aeruginosa secretome and is an important virulent factor [23]. Moreover, porcine elastase
Int. J. Mol. Sci. 2022, 23, 15253 8 of 23
was commercially available to perform some preliminary degradation tests, so we de-
cided to focus on this protease for further studies.
A single copy of a lasB gene encoding elastase (LasB protein) was detected in the
genome of P. aeruginosa IIVV-SD1 strain (Supplementary material Table S2 for the nucle-
otide sequence of the lasB gene and the corresponding amino acid sequence of LasB pro-
tein). It encodes a protein of 498 amino acids with a molecular weight of 53,687 Da and a
5.99 isoelectric point. The corresponding LasB protein exhibits an amino acid identity
ranging between 98.59% and 100% (WP_058140091.1) with many different proteins anno-
tated as M4 family elastase LasB proteins from other P. aeruginosa strains.
2.7. In Vitro Silk Biodegradation by Commercial Enzymes
Both commercial enzymes, porcine elastase and subtilisin A from B. licheniformis,
were able to digest raw silk when incubated at 37 °C for 24 h, producing a significant
release of amino acids in the reaction medium (Figure 5). Clear and visually evident deg-
radation of the silk ball was also detected in the reaction tubes at the end of the incubation.
In fact, the silk ball disappeared almost completely and only a few small, isolated fila-
ments could be observed as a result of the degradative process. Degradation rate was de-
pendent on enzyme concentration and for the highest concentrations (50 and 100 μg/mL)
we observed that subtilisin produced a greater silk degradation than elastase, as evident
from the higher amount of L-Leucine equivalents released into the reaction solution.
Figure 5. L-Leucine equivalent (mM) release after incubation (37 °C/24 h) of raw silk with different
amounts of the commercial protease porcine elastase and B. licheniformis subtilisin A. The results
shown are the average of two different experiments performed in triplicate.
2.8. Heterologous Expression of a P. aeruginosa LasB Elastase-Encoding Gene in E. coli and Its
Role in Silk Biodegradation
A recombinant LasB-SUMO protein was efficiently expressed in E. coli using pET
SUMO protein expression system (Figure 6A). Despite the fact that this system was de-
signed to produce soluble fusion proteins with a small ubiquitin-like modifier (SUMO) to
allow expression, purification, and generation of soluble recombinant proteins in E. coli,
in our case the recombinant protein was insoluble and was located in inclusion bodies. In
order to try to increase the production of soluble recombinant proteins, different modifi-
cations were attempted, such as induction using lower concentrations of IPTG, growth at
Protease (g/mL)
L-Leucine equivalent (mM)
0
2
4
6
8
10
Porcine elastase
Subtilisin A
Int. J. Mol. Sci. 2022, 23, 15253 9 of 23
30 °C, or growth in a poorer culture medium or without agitation. However, all tests at-
tempting to increase the solubility of the recombinant protein were unsuccessful. For that
reason, we had to solubilize the inclusion bodies using high concentrations of guanidine
hydrochloride. Once solubilized, the recombinant protein was purified on a Ni-NTA resin
and directly refolded, attached to the resin using successive washing steps with buffers
containing decreasing amounts of guanidine hydrochloride until a final wash with a
guanidine hydrochloride-free refolding buffer (Figure 6A).
Figure 6. (A) Expression in E. coli and purification of SUMO-LasB recombinant protein by Ni-NTA
affinity chromatography. Lanes: recombinant SUMO-CAT protein bound to Ni-NTA resin (1); Spec-
tra multicolor Broad Range Protein Ladder (Thermo Fisher Scientific, Waltham, MA, USA) (2); re-
combinant SUMO-LasB protein bound to Ni-NTA after refolding (3). (B) Raw silk enzymatic deg-
radation by SUMO-LasB recombinant protein as compared with different negative controls (C-) in-
cluding incubation of silk in the presence of a recombinant SUMO-CAT protein attached to a Ni-
NTA resin. The results shown are the average of two different experiments performed in triplicate.
The refolded recombinant protein had an approximate molecular weight of 67 kDa
as estimated by SDS-PAGE gel which corresponds fairly closely to the sum of the molec-
ular weights of SUMO (13 kDa) and LasB (53.687 kDa) (Figure 6A; lane 3). The incubation
of raw silk with the refolded recombinant SUMO-LasB protein produced the release of L-
Leucine equivalent into the reaction medium, with the release being most evident at 6 and
12 h of incubation (Figure 6B). Different negative controls were performed in parallel to
ensure that the release of amino acids was due to enzymatic action of the recombinant
protein on the silk (Figure 6B). As it can be seen in Figure 6A, lane 3, after the refolding of
the recombinant protein some bands of smaller molecular size were detected in the gel.
These bands could correspond to fragments obtained by the partial degradation of the
recombinant protein. In fact, we cannot rule out that in the samples analyzed there may
be traces of proteases responsible for this partial degradation. The release of a small
amount of L-Leucine equivalent observed in the negative control SUMO-LasB protein
(Figure 6B), which tends to increase with incubation time, could be due to the existence of
the previously mentioned proteases.
Int. J. Mol. Sci. 2022, 23, 15253 10 of 23
2.9. Overexpression of a B. licheniformis Subtilisin Carlsberg-Encoding Gene Enhances
Silk Biodegradation
Although a similar strategy to that employed for the P. aeruginosa LasB protein was
tried for expression of a recombinant SUMO-subtilisin protein, all attempts were unsuc-
cessful. Therefore, a different strategy was chosen which consisted in the homologous ex-
pression in B. licheniformis IIVV-SD3 strain of a subC gene, encoding a subtilisin Carlsberg
isolated from that same strain.
Two different transformants (B1 and B2) carrying the recombinant plasmid
pHY300PLK-SubC, which includes a copy of the subC gene under the control of its own
promoter region, were analyzed to determine its number of copies. Thus, the amplifica-
tion specificity of subC and rpoB genes was checked by melting curve analysis (Supple-
mentary material. Figure S2A). The subC primer set showed a sharp peak for the PCR
product of the quantitative standard sample at 84.38 ± 0.02 °C. A single melting peak at
the same melting temperature was produced for the PCR product of the total DNA sam-
ple. The rpoB primer set showed a single melting peak for the quantitative standard as
well as for the total DNA sample at the same temperature of 77.58 ±0.02 °C. Every PCR
product also yielded prominent bands with expected sizes of 100 and 70 bp, respectively,
after gel electrophoresis analysis. The identity of amplified products was additionally con-
firmed by DNA sequencing. These results indicated that non-specific PCR products were
not detected in the analyzed temperature range with the primer sets used (Supplementary
material. Figure S2A). The ratio between mass and copy number was calculated for each
PCR product resulting in 1 ng corresponding to 9.26 × 109 copies for subC PCR product
and 1.32 × 1010 copies for rpoB PCR product. The standard curves for subC and rpoB each
ranged from 1 × 106 to 1×1010 copies (Supplementary material. Figure S2B) obtained. Both
curves were linear in the range tested (R2 > 0.998) in the triplicate reactions. The slopes of
the standard curves for subC and rpoB were −3.755 and−3.061, respectively, with an am-
plification efficiency of 84.1% for subC and 107.6% rpoB. Standard curves were used for
relative quantification. For relative quantification, subC and rpoB were used as the target
and reference genes, respectively, and the plasmid copy number was determined by the
2−ΔΔCT calculation (Supplementary material. Figure S2C). Since B. licheniformis only has one
copy of the rpoB gene in its genome, the relative quantification showed that both B. lichen-
iformis IIVV-SD3 transformants analyzed harboring the pHY300PLK-subC plasmid con-
tained, in the middle of the exponential phase, an average of 7 more copies of the subC
gene than the B. licheniformis IIVV-SD3 wild type and accordingly, the average plasmid
copy number in each transformant was 7.
As seen in Figure 7A both transformants exhibited a delayed growth rate as com-
pared to that of B. licheniformis growth in the presence of raw silk. This delay in growth
could be due, among other factors, to the presence of the antibiotic tetracycline added to
the culture medium as a selective pressure to favor the maintenance of the plasmid carry-
ing the additional copies of the subC gene, or some growth interference due to the plas-
mid.
Int. J. Mol. Sci. 2022, 23, 15253 11 of 23
Figure 7. Stimulation of raw silk biodegradation in B. licheniformis B1 and B2 transformants carrying
8 copies (1 chromosomal copy and 7 plasmid copies) of the subC gene compared to the biodegrada-
tion rate observed in the wild-type strain IIVV-SD3 carrying a single chromosomal copy of the subC
gene. (A) Growth rate estimated as viable cells or cfu/mL. (B) Release of L-Leucine equivalent (μmol)
into the culture media from silk biodegradation. Given the different growth rates observed in the
transformants compared to the wild-type IIVV-SD3 strain, the results were normalized to the num-
ber of viable cells (cfu) detected in the cultures and expressed as CGUs (Cellular Growth Units).
One CGU equals 1 × 107 viable cells. The results shown are the average of two different experiments
performed in triplicate.
Therefore, in order to be able to compare the results of amino acid release into the
culture medium from silk biodegradation, the normalized results are shown in Figure 7B,
as a function of the number of viable cells present at each moment in the culture. Thus we
can see how, with equal numbers of cells, both B1 and B2 transformants, carrying a total
of 8 copies of the subC gene (1 chromosomal copy and 7 plasmid copies), showed an en-
hanced silk biodegradation capacity between days 2 and 6 of the experiment. More pre-
cisely, by day 4 of culture, transformant B1 exhibited a biodegradative capacity (estimated
as a function of the levels of amino acids present in the culture supernatant) that was 7.36
times higher than that observed in the wild-type strain. In the case of transformant B2, the
biodegradative capacity was 7.20 times higher. This result confirmed that the trans-
formants carrying additional copies of the subC gene were able to perform more efficient
silk biodegradation compared to the wild-type strain. At the end of the experiment,
growth of both transformants in the presence of silk was 1.63–1.95 times greater than that
observed in the wild type strain, most likely due to utilization of the released amino acids
as a source of carbon, nitrogen and energy.
3. Discussion
Despite the importance of the pine processionary as a forest pest and the significant
allergic reactions it causes, there are few studies on its raw silk. To our knowledge only
Shaw and Smith [11] analyzed pine processionary silk in an old work to conclude that its
fibroin had some unusual properties including a higher content of amino acids with hy-
drophilic side-chains, and also an increased resistance to acid hydrolysis as compared to
silk from Tiger moth (Arctia caja) and Oak Eggar (Lasiocampa quercus). Unfortunately, these
authors did not use silkworm silk, which is certainly the most studied, in their study.
According to our work, there are many similarities between pine processionary and
silkworm silk. Electron microscopy analysis shows that pine processionary silk has a very
similar macrostructure compared to that of silkworm silk. Thus, as in the case of silkworm
Int. J. Mol. Sci. 2022, 23, 15253 12 of 23
silk, each fiber contains 2 fibroin filaments coated with an outer glue-like coating, proba-
bly made of sericin [1]. Amino acid analysis of raw pine processionary silk indicated a
predominance of non-polar hydrophobic amino acids such as glycine, alanine, valine, leu-
cine, and tyrosine, and polar hydrophilic amino acids including neutral (serine) and acidic
(aspartic and glutamic acid) amino acids. This amino acid composition was very similar
to that observed by Shaw and Smith [11] who reported that the 3 most abundant amino
acids after hydrolysis of pure fibroin were glycine, serine, and alanine, exactly the same
ones that appear in our analysis. Small differences in amino acid composition could be
due to the different analytical methods used and also to the fact that whereas Shaw and
Smith analyzed pure fibroin, we analyzed raw silk (sericin plus fibroin). In a different
study, Do and colleagues [24] reported that the most abundant amino acids in silkworm
fibroin pretreated to remove sericin and other impurities were glycine (40.4%), alanine
(30.1%) and serine (10.2%). More recently Bungthong and colleagues [25] analyzed the
amino acid profile of silkworm silk after water and enzyme extraction, and although the
methodology is totally different and therefore the results cannot be easily compared, again
glycine was the most abundant amino acid with both serine and alanine also particularly
abundant. Therefore, we can conclude that pine processionary silk and silkworm silk have
a very similar amino acid composition, with a predominance of non-polar hydrophobic
amino acids (glycine, alanine) and a neutral hydrophilic amino acid like serine.
Since silk is one of the most abundant naturally derived protein polymers it should
come as no surprise that there are numerous microorganisms and mechanisms involved
in its biodegradation. The isolation from silk nests in an advanced state of degradation of
B. licheniformis and P. aeruginosa strains with the ability to biodegrade silk confirmed this
fact. The biodegradative capacity was confirmed by studies with pure cultures in minimal
liquid medium indicating that these microorganisms can use the peptides and amino ac-
ids released in the degradation process as a source of carbon, nitrogen, or energy. Analysis
by both optic and electron microscopy also confirmed that these microorganisms are able
to degrade silk to the point of completely destroying the native structure of the filaments.
The biodegradative capability of B. licheniformis, in the experimental conditions tested,
was higher than that observed for P. aeruginosa, as deduced from the higher concentration
of L-Leucine equivalent released into the culture media, even at the same or a lower num-
ber of cells. This higher biodegradative capacity could also be visually verified since B.
licheniformis cultures did not leave any silk filament remnants in liquid cultures, while
small filaments could be observed in the case of P. aeruginosa, which also suggested a
lower biodegradative capacity of this species. The higher biodegradation capability of B.
licheniformis compared to that observed by P. aeruginosa could be due to the different cleav-
age specificity of subtilisin and elastase proteins. In fact, and according this different
cleavage specificity, a much larger number of potential cleavage sites for subtilisin should
exist in pine processionary fibroin since this protease catalyzing the hydrolysis of proteins
and peptide amides shows high specificity for aromatic and hydrophobic amino acids at
the P1 position of the substrate [19,20]. These amino acids are particularly abundant in
pine processionary fibroin according to Shaw and Smith [11], and our own results.
Microbial degradation of silkworm silk by both bacterial and fungal strains had been
previously reported [10]. Regarding bacterial degradation, Seves and colleagues [12] iso-
lated a Burkholderia cepacia strain able to use fibroin as a sole source of carbon and nitrogen
for growth. A Variovorax paradoxus strain was also able to grow in a minimal medium
containing silk fibroin as the sole source of carbon and nitrogen [13]. A silk-degrading
enzyme, called fibroinase, was partially purified. The enzyme had a 21 kDA molecular
weight and was also able to degrade casein and, to a smaller extent, collagen and albumin,
although more accurate identification of this enzyme was not achieved. More recently B.
subtilis and P. fluorescens were isolated by their ability to degrade sericin [26].
We must not forget that silk fibroin from silkworm is of great interest as a biomaterial
with numerous applications in biomedicine including the development of films, mem-
branes, gels, sponges, powders, and scaffolds [27]. Applications include burn-wound
Int. J. Mol. Sci. 2022, 23, 15253 13 of 23
dressings, enzyme immobilization matrices, nets, vascular prostheses, and structural im-
plants [27], including nerve grafts in neuroscience [28]. In recent years, with an improved
understanding of the fundamental structures and properties of silk, along with options to
improve the purification of the native fiber structural core (fibroin) without residual con-
taminating proteins (e.g., sericin), degradable silk biomaterials have been generated
which are also biocompatible and show acceptable rates of biodegradability [29]. In this
context, it is not surprising that there are numerous studies analyzing the in vitro enzy-
matic degradation of fibroin and raw silk. Indeed, silk fibroin degrades in vitro and in
vivo in response to different proteolytic enzymes including actinase (a Streptomyces sp.,
enzymatic mix degrading actin) [30], α-chymotrypsin, collagenase, papain [27,29], prote-
ase XIV, and protease XXI (both proteases are really a mix of proteolytic enzymes isolated
from Streptomyces species including at least 3 different proteases, of which at least one is
an extracellular serine protease) [16].
In our case, and in order to identify some of the putative enzymes involved in raw
silk biodegradation, we discarded a classical approach consisting of cell fractionation and
successive purification steps using classical chromatographic techniques due to their la-
borious and time-consuming nature. Instead, we decided to adopt a strategy that takes
advantage of the enormous potential of genome sequencing techniques and genome anal-
ysis. Thus, sequencing of the B. licheniformis genome and analysis of the different prote-
ases and peptidases encoded in it allowed us to detect a total of 135 genes encoding en-
zymes with possible proteolytic activity (proteases and peptidases). This analysis com-
bined with a prediction of their putative extracellular location (extracellular enzymes) by
in silico analysis allowed us to conclude that a total of 20 proteases/peptidases could be
considered as secreted enzymes with an extracellular location. This group should include
the best candidates for enzymes involved in silk biodegradation, a process which, based
on the different microscopic observations made and the structure of silk itself, should be
extracellular. In fact, some evidence indicates that enzymatic degradation of silk and other
biomaterials is a two-step process. The first step is adsorption of the enzyme onto the sur-
face of the substrate through surface-binding domains, and the second step is hydrolysis
of the peptide bonds [30].
Analysis of these 20 extracellular proteases/peptidases allowed us to discard some of
them according to their biological function (e.g., those enzymes of the carboxypeptidase
type that are involved in wall peptidoglycan processing). Among the remaining enzymes,
subtilisin Carlsberg (a Ser-endopeptidase) particularly attracted our attention because it
is an extracellular alkaline serine protease, catalyzing hydrolysis of proteins and peptide
amides, that shows high specificity for aromatic and hydrophobic amino acids at the P1
position of the substrate [19,20]. This enzyme was, therefore, an ideal candidate for exten-
sive biodegradation of raw silk, as both sericin and fibroin would hypothetically contain
numerous potential cleavage sites. First analyses with commercial subtilisin from B. li-
cheniformis confirmed this fact: incubation of raw silk with the enzyme produced both a
significant release of amino acids into the reaction medium, and also a visually evident
degradation of the silk ball in the reaction tubes. Homologous overexpression of the lasB
gene encoding subtilisin Carlsberg in the strain from which it was isolated (strain IIVV-
SD3) confirmed its key role in silk biodegradation, as the transformants tested showed
higher degradation efficiency compared to the wild-type strain. Although in vitro biodeg-
radation data with commercial subtilisin confirmed its predominant role in silk biodegra-
dation, we cannot rule out that other proteases could be involved in the process. Among
the best candidates we should mention the two bacillopeptidases F detected (proteins 2
and 3 in Table S1—Supplementary Material). Bacillopeptidases F are extracellular fibrino-
lytic serine proteases [31] belonging to the superfamily of subtilisin-like serine proteases.
These enzymes are expressed at the beginning of the stationary phase in B. subtilis. Inac-
tivation of their encoding genes does not affect either vegetative growth or sporulation,
and they probably function as scavenging enzymes to supply the cell with amino acids
derived from protein degradation [32].
Int. J. Mol. Sci. 2022, 23, 15253 14 of 23
In a similar approach, analysis of the P. aeruginosa IIVV-SD1 strain sequenced ge-
nome allowed us to detect 9 extracellular proteases/peptidases. Among all of them, our
attention was primarily focused on elastase, or LasB protein, for several reasons. First,
because LasB is a protease with broad specificity, although it favors hydrophobic or aro-
matic amino acid residues, with preference to Phe and Leu at the P1′ and Gly in P1 posi-
tions [22]. Since Gly is higly abundant, and to a lesser extent Leu and Phe, in pine proces-
sionary fibroin [11], we could assume the existence of numerous cleavage sites for this
protease. Second, P. aeruginosa elastase, or pseudolysin, is the predominant protease in the
P. aeruginosa secretome and is an important virulence factor [23]. Finally, porcine elastase
was commercially available for performing some preliminary in vitro degradation tests,
although as we have already indicated above its cleavage specificity is different to that of
P. aeruginosa elastase. These tests confirmed that porcine elastase was able to efficiently
degrade raw silk. However, as there might be some differences in specificity, or degrada-
tion efficiency, between porcine and P. aeruginosa elastase, the latter was efficiently ex-
pressed in E. coli. The recombinant protein obtained was able to degrade crude silk in
vitro, as evidenced by its degradation into L-Leucine equivalent released into the reaction
medium.
Confirmation that P. aeruginosa elastase could efficiently degrade silk may have im-
portant biosanitary implications. P. aeruginosa is an opportunistic bacterial pathogen pre-
sent in many environments which can cause fatal and debilitating disease, especially in
patients whose immune responses are compromised, and who are unable to clear an ini-
tial infection [23]. In fact, P. aeruginosa can adapt to many situations, often adopting a bio-
film mode of growth within which the organism is able to survive antibiotic challenges
[33]. Since silkworm silk is the starting material for the production of numerous materials
for medical use in patients (sutures, films, membranes, gels, sponges, powders, and scaf-
folds, among others), as we mentioned above, potential colonization of P. aeruginosa of
these materials and their putative degradation by elastase should not be underestimated
and could be the cause of numerous problems in patients.
The finding that enzymes such as subtilisin and elastase can efficiently degrade pine
processionary silk, and possibly also silkworm silk (to our knowledge their ability to de-
grade silkworm silk has not been described so far) opens interesting perspectives on this
topic and expands the range of enzymes and microorganisms involved in the biodegra-
dation of silk materials.
Finally, we would like to emphasize some other practical applications that could be
derived from the findings described in this manuscript. First, we should mention that cur-
rent management of pine processionary moths includes a combination of preventive tech-
niques, such as planting policies and methods for early detection, curative methods such
as trapping of adults and larvae, elimination of winter nests, application of insecticides
and bioinsecticides, and pheromone treatment, especially in small urban areas [4]. Unfor-
tunately, these methods may provide insufficient levels of control or endanger the health
of human and domestic animals, particularly in urban parks and recreational suburban
areas. Moreover, insecticide applications in the anthropized sites may be ineffective, as
some parts of the plants remain untreated and spraying in inhabited areas often triggers
complaints from residents. Biological control has to be stringently applied to achieve a
satisfactory level of management [4]. However, in big forests, spraying with synthetic pes-
ticides, biopesticides such as Spinosad [34], Bt protein [35], or biocontrol agents like Bacil-
lus thuringiensis or entomopathogenic fungi [36,37] is the only method yielding some ef-
fectiveness [34]. Given the high protection to environmental conditions that silk nests pro-
vide to caterpillars, their disorganization or biodegradation by the application of strains
of subtilisin- or elastase-producing microorganisms could contribute to increasing the ef-
ficacy of other treatments. Our findings especially support the potential use of bioinsecti-
cide-producing bacterial strains with the ability to synthesize elastase or subtilisin to con-
trol pine processionary pest. Unfortunately, and to the best of our knowledge neither B.
licheniformis nor P. aeruginosa have known bioinsecticidal activity. Moreover, the use of
Int. J. Mol. Sci. 2022, 23, 15253 15 of 23
the latter microorganism as a BCA would not be possible given its character as an im-
portant opportunistic pathogen for humans and other animals [22]. However, it is inter-
esting to mention that some strains of B. thuringiensis include genes encoding for subtil-
isins in their genome [38]. In these strains subtilisin is a virulence factor that could be
involved in efficient larval body utilization during the infection process [38]. It has also
been reported that proteases, including subtilisin, play a significant role in the conversion
of δ-protoxins to active toxins [39,40]. Putting together all these data we could hypothesize
that the selection of B. thuringiensis strains with both good subtilisin and bioinsecticidal
activities toward pine processionary caterpillars could be a more efficient treatment than
biofumigation with Bt protein alone, since the larvicidal effect of the endotoxin would be
added to the biodegradative effect on the silk. The inoculation by biofumigation of these
kind of selected strains on silk nests could allow their active growth in the nests and thus
favor a more active transmission by a higher rate of infection among caterpillars probably
allowing an improved control of the pest.
Finally, as we have indicated above, in some countries the removal and subsequent
burning of winter nests is a common management practice to reduce the incidence of the
pest [4,18]. As we mentioned, the amino acids alanine, glycine and serine are particularly
abundant in silk, and to a lesser extent other amino acids such as aspartic, glutamine, ty-
rosine and arginine are also present. All these amino acids have many different industrial
applications and are obtained by chemical or biological methods (fermentations carried
out by selected microorganisms) [41]. It is possible that biodegradation of pine proces-
sionary nests under controlled conditions in an industrial environment, especially by B.
licheniformis, which has a greater biodegradation capacity than P. aeruginosa, could be an
interesting alternative for the industrial production of some of these amino acids. The case
of serine is particularly interesting since this amino acid is currently used in industries
ranging from food to cosmetics, and demonstrates many novel potential applications in
pharmaceutical industries [42]. Currently, L-serine production relies mainly on extraction
from protein hydrolysates, chemical synthesis, or enzyme or cellular conversion from the
precursor glycine plus a C1 compound such as methanol. However, dependence on high-
priced substrates, low yield, and environmental pollution make current production meth-
ods less attractive and non-sustainable [42]. Perhaps production of serine from the bio-
degradation of a waste material, such as silk from pine processionary nests, could be an
interesting alternative worth investigating, although is evident that some kind of exopep-
tidase should also be involved in the degradation of the peptides produced by the subtil-
isin activity in order to release serine and allow its accumulation into the culture super-
natant.
In summary, we would like to highlight that the finding that silk from pine proces-
sionary, and possibly also silkworm silk, can be biodegraded by subtilisin and elastase
type enzymes opens up a wide range of perspectives that deserve to be investigated, in-
cluding their repercussions in biomedicine, pest control and possible applications as a re-
newable source for the industrial production of certain amino acids.
4. Materials and Methods
4.1. Silk Processing
Silk was obtained from nests collected from a pine forest in the area known as La
Candamia (42°34′54.6″ N 5°32′08.0″ W), close to the city of León (Spain). For the isolation
of putative silk degrading microorganisms small silk samples of 1–2 g were taken in the
field directly from nests with an evident state of degradation using sterile scissors and
forceps and immediately introduced into sterile Falcon tubes to avoid any accidental con-
tamination. The silk used for further microbial or enzymatic degradation studies in the
laboratory was obtained from the collection of young nests that were placed in plastic
bags, and autoclaved to kill any caterpillars. The silk was then manually cleaned using
combs and tweezers in order to remove any remains of sticks, pine needles, caterpillars
Int. J. Mol. Sci. 2022, 23, 15253 16 of 23
and caterpillar excrement naturally present in the nests. After careful cleaning, the silk
was re-sterilized in an autoclave and stored in sterile Falcon tubes at 4 °C until use.
4.2. Acid Hydrolysis of Silk and Amino Acid Analysis by HPLC
Acid hydrolysis of raw silk was carried out following the methodology described by
Hess and colleagues [43] with minor modifications. Briefly, 2 mg silk was suspended in 1
mL 6N HCl in a glass tube. Phenol (0.02%) was included as a protective agent to reduce
the loss of some residues during hydrolysis [44]. Traces of oxygen were replaced by re-
peated flushing with nitrogen. The closed tube was heated at 110 °C for 24 h to achieve
complete hydrolysis. Next, the liquid was evaporated in a rotary evaporator with a vac-
uum pump(Buchi R300, Barcelona, Spain) at 65 °C for 8 h. The residue was dissolved in
water (1 mL) and evaporated to remove further HCl. The final residue was dissolved in
200 μL of MQ water and stored at −20 °C until use. The amino acid composition of the
hydrolyzate was analyzed, previous o-phthalaldehyde derivatization, by reversed-phase
HPLC (RP-HPLC) [45] on a Zorbax Eclipse XDB C18 (4.6 × 150 mm; 5 μm) column (Agilent
Technologies, Saint Claire, CA, USA) using an Agilent 1200 Liquid Chromatograph
equipped with a quaternary pump delivery system (Saint Claire, CA, USA) (G1311A), a
preparative autosampler (G1329A), a diode array multi-wavelength detector (G7115A), a
fluorescence detector (G1321A), and an analytical fraction collector (G1364F) equipped
with an Autosampler Thermostat (G1330B). Samples of 2.5 μL were injected, and detec-
tion was carried out at 338 nm. Chromatographic conditions were the same as reported
by Bartolomeo and Maisano [45]. Quantification of amino acids was carried out according
to the area of their respective peaks by comparison with authentic amino acid peaks.
Amino acid standard samples contained 20, 50, 130, 250, or 500 pmol/μL of an amino acid
standard mixture together with 0.5 mM norvaline.
4.3. Isolation and Identification of Silk-Degrading Bacterial Strains
Twenty mg of silk collected from an old nest with evident symptoms of biodeterio-
ration were deposited at the bottom of a 10 mL polypropylene sterile tube. Next, 2.0 mL
of yeast extract (0.25%, w/v), and 0.1 mL of a salt mixture [ZnSO4 x 7H2O, 4 g/L; FeSO4 x
7H2O, 9 g/L; CuSO4 x 7H2O, 0.18 g/L; H3BO3, 0.026 g/L; (NH4)3Mo4O7 x 4H2O, 0.017 g/L,
MnSO4 x 4H2O] were added and the mix was brought to a final volume of 5 mL with
sterile water. The mix was incubated up to 60 days at RT. Once silk degradation was vis-
ually observed, two more enrichment rounds were carried out under the same conditions,
except 0.1% glucose was added to accelerate bacterial growth, and in turn, the biodegra-
dation process. Then tenfold dilutions were spread on the surface of Plate Count Agar
(PCA) (Condalab, Torrejón de Ardoz, Spain) plates containing natamycin (200 μg/mL) to
avoid fungal growth. Plates were incubated at 28 °C until bacterial colonies developed.
Bacteria representative of different macro and microscopic morphological types were se-
lected at random and conserved in Nutrient Agar plates (Condalab) at 4 °C until use.
To test which of the bacterial isolates had the ability to biodegrade silk, pure cultures
in the presence of sterile silk were developed as previously indicated, including 0.1% glu-
cose. Cultures were inoculated with 500 μL of a pure bacterial culture (OD 1 at 600 nm).
Positive biodegrading strains were selected for their ability to visually solubilize silk and
for their ability to release ninhydrin positive products from the degraded silk. Amino acid
release was quantified with ninhydrin as reported by Sun et al. [46] and the color pro-
duced was determined spectrophotometrically at 580 nm against a standard curve con-
taining known amounts of leucine.
Silk degrading isolates were identified by 16S rRNA sequencing. Briefly, genomic
DNA extraction was performed as described by Hopwood et al. [47]. Amplification of 16S
rRNA genes was carried out using oligonucleotides 27F and 1492R [48]. Isolates were
identified by comparison with the corresponding sequences of type strains found on Ez
Int. J. Mol. Sci. 2022, 23, 15253 17 of 23
Taxon-e database [49] (http://www.ezbiocloud.net/eztaxon/identify accessed on 18 Febru-
ary 2022). Sequence alignment as well as phylogenetic trees were carried out using the
MEGA 6 software [50].
4.4. Sequencing and Genome Assembly of Silk Degrading Strains
Genomic DNA was sent to Macrogen (Seoul, South Korea) for Illumina sequencing.
Illumina TruSeq Nano DNA Kit was used to generate the Illumina library according to
the manufacturer’s specifications. Illumina sequencing was performed on a Novaseq-6000
producing paired-end 2 × 150 bp reads. Raw Illumina reads were analyzed with FASTQC
v0.11.8 (http://www.bioinformatics.babraham.ac.uk/projects/fastqc accessed on 18 Febru-
ary 2022) to obtain quality statistics, then Trimmomatic v0.38 [51] was used to remove
adapter sequences and trim out bases of low quality (minimum base quality 35 and min-
imum read length 35 bp). The Illumina reads were then aligned against reference genomes
(B. licheniformis GCF_002074095.1 and P. aeruginosa GCF_000006765.1) with BWA v0.7.17
(http://bio-bwa.sourceforge.net/ accessed on 18 February 2022). During mapping, dupli-
cate reads were removed using Sambamba v0.6.8 (http://lomereiter.github.io/sambamba/
accessed on 18 February 2022). After removing duplicates and identifying variants with
SAMTools (http://samtools.sourceforge.net/ accessed on 18 February 2022) information
about each variant was gathered and classified by chromosomes or scaffolds. In order to
discover annotation information, such as amino acid changes by variants, SnpEff v4.3t
(http://snpeff.sourceforge.net/ accessed on 18 February 2022) was used.
Genomes of the B. licheniformis and P. aeruginosa strains were assembled using
SPAdes v. 3.15.4 [52]. Quality of the results was analyzed using Quast v. 5.2.0 [53], com-
paring it to the reference genome of each species to check its coverage. The assembled
genomes were processed with BUSCO v5.3.25 [54] to make a preliminary annotation and
predict the proteins with Prodigal [55]. Subsequently, the predicted proteins were anno-
tated with Blastp v2.9.0-2 [56] against the NCBI SwissProt database [57]. Finally, using an
R v4.2.1 [58] script, a functional annotation was obtained by propagating the annotation
to obtain the GO terms [59] and the InterPro [60] protein families corresponding to each
annotated protein.
Both whole genome shotgun projects were deposited at DDBJ/ENA/GenBank as a
Bioproject (ID PRJNA894474) that includes the B. licheniformis (IIVV-SD3 strain) genome
project (accession number JAPDGT000000000) and the P. aeruginosa (IIVV-SD1 strain) ge-
nome project (accession number JAPDGU000000000).
4.5. Silk Biodegradation in Liquid Media
Analysis of silk degradation in liquid media was carried out in triplicate in 50 mL
Falcon tubes with a perforated plug and a valve for allowing gas exchange. Tubes con-
tained 10–20 mg of sterile silk suspended in 30 mL of minimal medium [K2HPO4, 40 mM;
KH2PO4, 22 mM; MgSO4 x 7H2O, 0.8 mM; KNO3, 10 mM; glucose, 0.5% (w/v) and 30 μL of
a salt mixture as indicated above in Section 4.3]. Tubes were inoculated with 106 bacterial
cells/mL and incubated at 30 °C. Samples (500 μL) were removed at different times and
silk degradation quantified by measurement of free amino acids in the culture superna-
tant by performing a ninhydrin test [46]. Amino acid quantification was carried out at 570
nm by comparison with a standard curve of leucine.
4.6. Scanning Electron Microscopy (SEM) and Transmission Electron Microscopy (TEM)
Raw silk samples were processed according the Bertazzo protocol [61]. Briefly, silk
samples were fixed with electron microscopy grade glutaraldehyde (TAAB Laboratories,
Berks, UK) at a final concentration of 2.5% in phosphate buffered saline (PBS) for 2 h at 4
°C. Samples were washed three times in PBS. Next, they were refixed in the dark at RT
with 2% osmium tetroxide (TAAB Laboratories) in PBS for 2 h. Finally, three 30-min
Int. J. Mol. Sci. 2022, 23, 15253 18 of 23
washes were performed with PBS. Next, samples were dehydrated by immersion in solu-
tions of increasing ethanol concentration for 30 min each step (30%, 50%, 70%, 90%, 3 ×
96% and 3 × 100%). Sample drying was carried out using a critical-point desiccator model
CPD 030 from BAL-TEC Inc (Liechtenstein). Samples were immediately gold-coated and
mounted on aluminum stabs to be examined with a Scanning Electron Microscope model
JSM-6480 LV from JEOL (Tokyo, Japan).
Samples for TEM analysis were prepared according to the protocol described by
Wang et al. [28] and observed under a transmission electron microscope model JEM-1010
(JEOL).
4.7. Enzymatic Assays for In Vitro Silk Degradation
In vitro enzymatic degradation of silk was tested using commercial porcine pancre-
atic elastase (≥8 U/mg of protein) (Worthington Biochemical Corporation, Lakewood, NJ,
USA) or commercial lyophilized subtilisin A (7–15 U/mg of protein) from B. licheniformis
(Merck KGaA, Darmstadt, Germany). Reactions contained 2 mg clean, sterile silk and dif-
ferent amounts of enzyme and were carried out under the assay conditions indicated by
the supplier.
4.8. DNA Isolation and Manipulation
Small-scale total DNA from B licheniformis and P. aeruginosa was extracted using Il-
lustraTM Bacteria genomicPrep Mini Spin Kit (Illustra, GE Healthcare Bio-Sciences AB,
Uppsala, Sweden). Large-scale total DNA isolation was carried out as described by
Hopwood et al. [47]. Small scale plasmid DNA isolation from E. coli was carried out using
the boiling method [62] or alternatively using IllustraTM plasmidPrep Mini Spin Kit (Illus-
tra). DNA manipulations were performed according to standard procedures [63].
4.9. Heterologous Expression in E. coli of a P. aeruginosa lasB Elastase-Encoding Gene
A P. aeruginosa LasB-encoding gene was amplified from genomic DNA by PCR using
F-LasB (5′-ATGAAGAAGGTTTCTACGCTTGACC-3′) and R-LasB (5′-TTACAAC-
GCGCTCGGGCA-3′) primers and Phusion Hot Star II DNA polymerase (Thermo Fisher
Scienfic, Waltham, MA, USA). The PCR fragment was ligated to ChampionTM pET SUMO
expression vector (Thermo Fisher Scientific) and the ligation reaction was transformed
into E. coli One Shot® Mach1TM (Thermo Fisher Scientific) competent cells following the
manufacturer’s instructions. The resultant recombinant plasmid (pET SUMO-LasB) was
sequenced to confirm that no nucleotide changes had occurred during PCR amplification,
and the insert had been introduced in the right orientation.
Next, this recombinant plasmid was introduced into competent cells of E. coli
BL21(DE3) One Shot strain (Thermo Fisher Scientific) to check putative LasB overexpres-
sion. A selected recombinant clone was grown in an orbital shaker (37 °C/220 r.p.m) in LB
liquid medium (100 mL) containing 50 μg/mL of kanamycin until an O.D. 600 nm of 0.5
was reached. Overexpression was induced by adding 0.75 mM IPTG and incubating at 37
°C for an additional 6 h. Cells were recovered by centrifugation and pellet resuspended
in Buffer A (NaH2PO4, 100 mM; Tris-HCl, 10 mM; guanidine hydrochloride, 6M; NaCl, 0.5
M; 2-mercaptoethanol, 1 mM; pH 8.0). Cells were disrupted by sonication in an ice bath
using 15 cycles of 3–4 s bursts (90–100 W) with a Branson Sonifier B-12 (Thermo Fisher
Scientific). Inclusion bodies were solubilized at RT for 4 h in a rotating arm. Cellular debris
were removed by centrifugation (4 °C/15,000× g/10 min).
Recombinant protein was purified onto 0.5 mL of HisPureTM Ni-NTA resin (Thermo
Fisher Scientific), previously equilibrated by extensive washing in Buffer A, by overnight
incubation at 4 °C in a rotating arm. Contaminant and unbound proteins were removed
by washing (3 times) with 10 vol Buffer A containing 20 mM imidazole. Next the resin
was packed into a 1 mL small column and the recombinant protein was refolded and im-
mobilized on the Ni-NTA matrix using step-by-step removal of guanidine hydrochloride
Int. J. Mol. Sci. 2022, 23, 15253 19 of 23
using a Refolding Buffer (Tris-HCl, 50 mm; NaCl, 100 mM; glycerol, 10% (w/v); pH 7.0)
containing decreasing concentrations (5M, 4M, 3M, 2M, 1M, 0.5M, 0.25M) of guanidine
hydrochloride. All the refolding steps were done at 4 °C by slowly adding 2 mL of each
Refolding Buffer to the column. Finally, the protein bound to the Ni-NTA matrix was con-
served in Refolding Buffer at 4 °C until use. A small amount of resin (10–20 μL) was ana-
lyzed by SDS-PAGE to check the purified refolded protein. Protein concentration was vis-
ually estimated by comparison with known amounts of BSA run in an SDS-PAGE gel in
parallel.
4.10. Analysis of Silk Degradation by Recombinant Elastase
The ability of recombinant LasB-SUMO protein bond to Ni-NTA resin to degrade silk
was tested in in vitro reactions containing 10 μg recombinant protein and 2 mg silk in a
total volume of 1.5 mL Tris-HCl, 100 mM (pH 8.0) buffer. Reactions were incubated at 37
°C up to 12 h in a rotating arm. Samples of 100–200 μL were removed at different times
and silk degradation was estimated by release of amino acids into the solution using the
ninhydrin assay.
4.11. Homologous Expression in B. licheniformis of a Subtilisin Carlsberg-Encoding Gene (subC)
A recombinant plasmid pHY300PLK-pSubC was obtained by cloning the subtilisin
Carlsberg-encoding gene (subC) located in the genome of B. licheniformis IIVV-SD3 into
the shuttle vector pHY300PLK [64]. The gene and its promoter region were PCR-amplified
from genomic DNA using F_PSubC (5′-AATCAGGATCCCCGTTTCTGTATGCGATA-3′)
and R_PSubC (5′-CTTATCCCGGGTTATTGAGCGGCAGCTTCGAC) primers and high
affinity Phusion Hot Star II DNA polymerase (Thermo Fisher Scienfic). A PCR product of
the expected size was purified from an agarose gel using the Freeze-Squeeze method [65]
and later digested with BamHI and XbaI restriction enzymes in order to generate compat-
ible cohesive ends that allowed its cloning in the BamHI-XbaI digested pHY300PLK vec-
tor. Ligation mix was transformed into E. coli ET12567 strain competent cells following
standard protocols [63]. A clone containing the recombinant plasmid pHY300PLK-pSubC
was selected by restriction analysis and sequenced to confirm the absence of undesired
mutations. Next the recombinant pHY300PLK-pSubC plasmid was introduced into B. li-
cheniformis IIVV-SD3 by electroporation following the protocols described by Xue et al.
[66]. DNA from transformants was isolated with GeneJET Plasmid Miniprep Kit (Thermo
Fisher Scientific) and the presence of recombinant plasmid was checked by PCR using
pHY300PLK_F (5′-GTCAGATTTCGTGATGCTTGTC-3′) and pHY300PLK_R (5′-
GGATCAACTTTGGGAGAGAGTTC-3′) primers. These primers anneal to both sides of
the multicloning site (MCS) of the pHY300PLK vector and allowed amplification of the
whole insert cloned in pHY300PLK-pSubC plasmid, whereas no amplification was ob-
served from total DNA from B. licheniformis IIVV-SD3·used as negative control. Addi-
tional verification was carried out by restriction analysis of plasmid DNA isolated from
the transformants. Two transformants were selected for further studies and copy number
of subC gene determined by quantitative PCR (qPCR). Total DNA from both trans-
formants and IIVV-SD strain was isolated using Illustra™ Bacteria genomicPrep Mini
Spin Kit (Illustra).
For plasmid copy number determination in the transformants, two primer sets spe-
cific to subC and rpoB genes, this last encoding RNA polymerase β-subunit, were de-
signed. The subC gene was present as a single copy in pHY300PLK-pSubC plasmid and
also in the chromosomal DNA, and likewise the rpoB gene has only one copy in the bac-
terial genome. Primer sets were designed using Primer3 software [67]. A 100 bp internal
fragment of subC gene was amplified using F_subC-qPCR (5′-TTCACAAGTCCGCAAC-
CGTC-3′) and R_subC-qPCR (5′-TTATTGAGCGGCAGCTTCGAC-3′) primers. In a simi-
lar way a 70 bp internal fragment of rpoB gene was amplified using F_rpoB-qPCR (5′-
ACCTCTTCTTATCAGTGGTTTCTTGAT-3′) and R_rpoB-qPCR (5′-CCTCAATTGGCGA-
Int. J. Mol. Sci. 2022, 23, 15253 20 of 23
TATGTCTTG-3′) primers. Internal fragments of both genes were amplified by conven-
tional PCR. The reaction mixture of 50 μL contained 0.1 mM dNTP, 0.5 μM of each primer,
0.05 U Paq5000 DNA polymerase and 25 ng of B. licheniformis genomic DNA as a template.
PCR amplification was performed with a Mastercycler Gradient thermocycler (Eppen-
dorf, Hamburg, Germany), according to the following program: an initial denaturation at
98 °C for 3 min, followed by 30 cycles of 30 s at 98 °C, 30 s at 60 °C and 1 min at 72 °C; a
final extension at 72 °C for 10 min. Both PCR products were purified using the NucleoSpin
Gel and PCR Clean-up kit (Macherey-Nagel GmbH & Co. KG, Duren, Germany) and
quantified on NanoDrop 2000 (Thermo Fisher Scientific). Since the size and nucleotide
composition of each amplicon are known, the number of copies of each PCR amplified
fragment was easily estimated using an online calculator
(http://cels.uri.edu/gsc/cndna.html accessed on 3 October 2022). Then, PCR products were
serially 10-fold diluted and used as templates to generate qPCR standard curves, one for
each gene (Supplementary material. Figure S2A).
The B. licheniformis IIVV-SD3 wild type and two B. licheniformis IIVV-SD3 trans-
formants harboring pHY300PLK-pSubC plasmid were cultured in flasks containing 50 mL
of LB (supplemented with 30 μg/mL tetracycline for culture transformants). Total DNA
was extracted from both cultures when they reached approximately half the exponential
phase (OD600nm 0.8). DNA extraction was performed using the IllustraTM bacteria ge-
nomicPrep Mini Spin Kit (GE Healthcare), following manufacturer’s instructions. The
concentration of extracted DNA was measured using NanoDrop 2000 (Thermo Fisher Sci-
entific). Genomic DNA was serially diluted 10-fold and copy number of each gene was
estimated using the previous standard curves (Supplementary material. Figure S2B).
Real-time QPCR SYBR Green amplification and analyses were performed using a
Mx3005P QPCR system and software (Agilent technologies). The real time qPCR mixture
of 20 μL contained 1X SYBR Green master mix (Takara, Shiga, Japan), 0.2 μMol of each
primer and 5 μL bacterial DNA. Nuclease-free water was used to bring the reaction vol-
ume to 20 μL. The thermal cycling protocol was as follows: initial denaturation for 2 min
at 95 °C followed by 40 cycles of 30 s at 95 °C and 30 s at 60 °C. After amplification, a
melting curve analysis with a temperature gradient of 0.1 °C/s from 55 to 95 °C was per-
formed to confirm that only specific products were amplified. All analyses were per-
formed as three independent experiments with three replicates for each dilution.
5. Conclusions
Our current study extends the knowledge on the molecular mechanisms involved in
the microbial degradation of silk materials and fibroin-like proteins. For the first time we
demonstrate that bacteria such as P. aeruginosa and B. licheniformis can efficiently biode-
grade silk materials, and more specifically silk produced by the pine processionary. For
this purpose, these microorganisms use common enzymes in the bacterial world such as
elastase produced by P. aeruginosa or subtilisin produced by B. licheniformis. Therefore,
any microorganism producing these types of enzymes would be susceptible to degrade
silk materials.
Supplementary Materials: The following supporting information can be downloaded at:
https://www.mdpi.com/article/10.3390/ijms232315253/s1.
Author Contributions: Conceptualization, R.C. and J.J.R.C.; methodology, A.D.-G., R.C. and
J.J.R.C.; data curation, A.D.-G.; formal analysis, J.J.R.C.; investigation, A.D.-G., A.I., C.C.-P. and R.C.;
writing—original draft, R.C. and J.J.R.C.; writing—review and editing, R.C. and J.J.R.C.; funding
acquisition J.J.R.C. All authors have read and agreed to the published version of the manuscript.
Funding: Alba Diez-Galán (EDU/556/2019), Ana Ibañez (EDU/529/2017) and Carla Calvo-Peña
(EDU/601/2020) were supported by a predoctoral contract from the Junta de Castilla y León and the
European Social Fund..
Institutional Review Board Statement: Not applicable.
Int. J. Mol. Sci. 2022, 23, 15253 21 of 23
Informed Consent Statement: Not applicable.
Data Availability Statement: Not applicable.
Acknowledgments: We thank Ulrich Schwaneberg (RWTH Aachen University, Germany) who pro-
vided us with the pHY300PLK plasmid for their kind support with the bioinformatics analysis.
Conflicts of Interest: The authors declare no conflict of interest.
References
1. Phan Nguyen, T.; Vinh Nguyen, Q.; Nguyen, V.-H.; Le, T.-H.; Quynh Nga Huynh, V.; Vo, D.-V.N.; Thang Trinh, Q.; Young Kim,
S.; Van Le, Q. Silk Fibroin-Based Biomaterials for Biomedical Applications: A Review. Polymers 2019, 11, 1933.
https://doi.org/10.3390/polym11121933.
2. Kaplan, D.; Adams, W.W.; Farmer, B.; Viney, C. Silk : Biology, Structure, Properties, and Genetics. In Silk Polymers; ACS Sym-
posium Series; American Chemical Society: Washington, DC, USA, 1994; pp. 2–16.
3. Pimentel, C.; Ferreira, C.; Nilsson, J.Å. Latitudinal gradients and the shaping of life-history traits in a gregarious caterpillar. Biol.
J. Linn. Soc. 2010, 100, 224–236. https://doi.org/10.1111/J.1095-8312.2010.01413.X.
4. Trematerra, P.; Colacci, M.; Athanassiou, C.G.; Kavallieratos, N.G.; Rumbos, C.I.; Boukouvala, M.C.; Nikolaidou, A.J.; Kontodi-
mas, D.C.; Benavent-Fernández, E.; Gálvez-Settier, S. Evaluation of Mating Disruption for the Control of Thaumetopoea
pityocampa (Lepidoptera: Thaumetopoeidae) in Suburban Recreational Areas in Italy and Greece. J. Econ. Entomol. 2019, 112,
2229–2235. https://doi.org/10.1093/jee/toz129.
5. Battisti, A.; Stastny, M.; Netherer, S.; Robinet, C.; Schopf, A.; Roques, A.; Larsson, S. Expansion of geographic range in the pine
processionary moth caused by increased winter temperatures. Ecol. Appl. 2005, 15, 2084–2096. https://doi.org/10.1890/04-1903.
6. Galip, N.; Şanlıdağ, B.; Babayiğit, A.; Bahçeciler, N.N. Cutaneous Allergic reactions to pine processionary caterpillar (Thaumet-
opoea Pityocampa): A complicated cutaneous reaction in an infant and review of the literature. Turk. J. Pediatr. 2022, 64, 389–
393. https://doi.org/10.24953/turkjped.2021.385.
7. Kanat, M.; Alma, M.H.; Sivrikaya, F. Effect of defoliation by Thaumetopoea pityocampa (Den. & Schiff.) (Lepidoptera: Thaumeto-
poeidae) on annual diameter increment of Pinus brutia Ten. in Turkey. Ann. For. Sci. 2005, 62, 91–94. https://doi.org/10.1051/for-
est.
8. Liu, X.; Zhang, K.-Q.; Liu, X.; Zhang, K.-Q. Silk Fiber—Molecular Formation Mechanism, Structure-Property Relationship and
Advanced Applications. In Oligomerization of Chemical and Biological Compounds; IntechOpen: London, UK, 2014; ISBN 978-953-
51-1617-2.
9. Reddy, N.; Yang, Y. Introduction to Natural Protein Fibers. In Innovative Biofibers from Renewable Resources; Springer: Berlin/
Heidelberg, Germany, 2015; pp. 157–158. https://doi.org/10.1007/978-3-662-45136-6_34.
10. Gutarowska, B.; Michalski, A. Microbial Degradation of Woven Fabrics and Protection Against Biodegradation. In Woven Fabrics;
Jeon, H.-Y., Ed.; InTech: London, UK, 2012; ISBN 978-953-51-0607-4.
11. Shaw, J.T.B.; Smith, S.G. Comparative studies of fibroins. III. The silk fibroin of the pine-processionary moth (Thaumetopoea
pityocampa)-An unusual β-protein. BBA Biochim. Biophys. Acta 1961, 46, 302–310. https://doi.org/10.1016/0006-3002(61)90753-3.
12. Seves, A.; Romanò, M.; Maifreni, T.; Sora, S.; Ciferri, O. The microbial degradation of silk: A laboratory investigation. Int. Bio-
deterior. Biodegrad. 1998, 42, 203–211. https://doi.org/10.1016/S0964-8305(98)00050-X.
13. Forlani, G.; Maria Seves, A.; Ciferri, O. A bacterial extracellular proteinase degrading silk fibroin. Int. Biodeterior. Biodegrad. 2000,
46, 271–275. https://doi.org/10.1016/S0964-8305(00)00099-8.
14. Nakanishi, T.; Yamamoto, T. Action and Specificity of a Streptomyces Alkalophilic Proteinase. Agric. Biol. Chem. 1974, 38, 2391–
2397.
15. De, S.; Chandra, A.L. Degradation of Organic Nitrogenous Streptomycete Wastes by a Soil. Folia Microbiol. 1979, 24, 473–477.
16. Horan, R.L.; Antle, K.; Collette, A.L.; Wang, Y.; Huang, J.; Moreau, J.E.; Volloch, V.; Kaplan, D.L.; Altman, G.H. In vitro degra-
dation of silk fibroin. Biomaterials 2005, 26, 3385–3393. https://doi.org/10.1016/j.biomaterials.2004.09.020.
17. Arai, T.; Freddi, G.; Innocenti, R.; Tsukada, M. Biodegradation of Bombyx mori silk fibroin fibers and films. J. Appl. Polym. Sci.
2004, 91, 2383–2390. https://doi.org/10.1002/app.13393.
18. Yiğit, Ş.; Akça, İ.; Bayhan, E.; Bayhan, S.; Tekin, F.; Saruhan, İ. Determining the Toxicity of Some Thyme Essential Oils Against
the Pine Processionary [Thaumetopoea pityocampa (Lepidoptera: Notodontidae)]. Atatürk Univ. J. Agric. Fac. 2019, 50, 226–230.
https://doi.org/10.17097/ataunizfd.518352.
19. Evans, K.L.; Crowder, J.; Miller, E.S. Subtilisins of Bacillus spp. hydrolyze keratin and allow growth on feathers. Can. J. Microbiol.
2000, 46, 1004–1011. https://doi.org/10.1139/w00-085.
20. Azrin, N.A.M.; Ali, M.S.M.; Rahman, R.N.Z.R.A.; Oslan, S.N.; Noor, N.D.M. Versatility of subtilisin: A review on structure,
characteristics, and applications. Biotechnol. Appl. Biochem. 2022, Volume, 1–18. https://doi.org/10.1002/bab.2309.
21. Kessler, E.; Safrin, M. Pseudomonas Elastinolytic and Proteolytic Enzymes. In Pseudomonas Methods and Protocols, Methods in
Molecular Biology; Filloux, A., Ramos, J.-L., Eds.; Springer Science Business Media: New York, NY, USA, 2014; Volume 1149, pp.
135–169, ISBN 9781493904723.
22. Dau, T.; Gupta, K.; Berger, I.; Rappsilber, J. Sequential Digestion with Trypsin and Elastase in Cross-Linking Mass Spectrometry.
Anal. Chem. 2019, 91, 4472–4478. https://doi.org/10.1021/acs.analchem.8b05222.
Int. J. Mol. Sci. 2022, 23, 15253 22 of 23
23. Everett, M.J.; Davies, D.T. Pseudomonas aeruginosa elastase (LasB) as a therapeutic target. Drug Discov. Today 2021, 26, 2108–
2123. https://doi.org/10.1016/j.drudis.2021.02.026.
24. Do, S.G.; Park, J.H.; Nam, H.; Kim, J.B.; Lee, J.Y.; Oh, Y.S.; Suh, J.G. Silk fibroin hydrolysate exerts an anti-diabetic effect by
increasing pancreatic β cell mass in C57BL/KsJ-db/db mice. J. Vet. Sci. 2012, 13, 339–344. https://doi.org/10.4142/jvs.2012.13.4.339.
25. Bungthong, C.; Wrigley, C.; Sonteera, T.; Siriamornpun, S. Amino acid profile and biological properties of silk cocoon as affected
by water and enzyme extraction. Molecules 2021, 26, 3455. https://doi.org/10.3390/molecules26113455.
26. Pandya, B.; Shetty, S. Bacterial degradation of sericin for degumming of silk fibers–A green approach. J. Appl. Biol. Biotechnol.
2021, 9, 89–95. https://doi.org/10.7324/JABB.2021.9513.
27. Cao, Y.; Wang, B. Biodegradation of Silk Biomaterials. Int. J. Mol. Sci. 2009, 10, 1514–1524. https://doi.org/10.3390/ijms10041514.
28. Wang, Y.; Liang, Y.; Huang, J.; Gao, Y.; Xu, Z.; Ni, X.; Yang, Y.; Yang, X.; Zhao, Y. Proteomic Analysis of Silk Fibroin Reveals
Diverse Biological Function of Different Degumming Processing From Different Origin. Front. Bioeng. Biotechnol. 2022, 9, 777320.
https://doi.org/10.3389/FBIOE.2021.777320.
29. Guo, C.; Li, C.; Vu, H.V.; Hanna, P.; Lechtig, A.; Qiu, Y.; Mu, X.; Ling, S.; Nazarian, A.; Lin, S.J.; et al. Thermoplastic moulding
of regenerated silk. Nat. Mater. 2020, 19, 102–108. https://doi.org/10.1038/s41563-019-0560-8.
30. Chen, K.; Umeda, Y.; Hirabayashi, K. Enzymatic hydrolysis of silk fibroin. J. Seric. Sci. Jpn 1995, 65, 131–133.
31. Meng, D.; Dai, M.; Xu, B.L.; Zhao, Z.S.; Liang, X.; Wang, M.; Tang, X.F.; Tang, B. Maturation of fibrinolytic bacillopeptidase F
involves both heteroand autocatalytic processes. Appl. Environ. Microbiol. 2016, 82, 318–327. https://doi.org/10.1128/AEM.02673-
15.
32. Wu, X.C.; Nathoo, S.; Pang, A.S.; Carne, T.; Wong, S.L. Cloning, genetic organization, and characterization of a structural gene
encoding bacillopeptidase F from Bacillus subtilis. J. Biol. Chem. 1990, 265, 6845–6850. https://doi.org/10.1016/S0021-
9258(19)39225-7.
33. Soares, A.; Alexandre, K.; Etienne, M. Tolerance and Persistence of Pseudomonas aeruginosa in Biofilms Exposed to Antibiotics:
Molecular Mechanisms, Antibiotic Strategies and Therapeutic Perspectives. Front. Microbiol. 2020, 11, 423.
https://doi.org/10.3389/fmicb.2020.02057.
34. Semiz, G.; Cetin, H.; Isik, K.; Yanikoglu, A. Effectiveness of a naturally derived insecticide, spinosad, against the pine proces-
sionary moth Thaumetopoea wilkinsoni Tams (Lepidoptera: Thaumetopoeidae) under laboratory conditions. Pest Manag. Sci. 2006,
62, 452–455. https://doi.org/10.1002/ps.1181.
35. Cebeci, H.H.; Oymen, R.T.; Acer, S. Control of pine processionary moth, Thaumetopoea pityocampa with Bacillus thuringiensis in
Antalya, Turkey. J. Environ. Biol. 2010, 31, 357–361.
36. Draganova, S.; Takov, D.; Pilarska, D.; Doychev, D.; Mirchev, P.; Georgiev, G. Fungal pathogens on some lepidopteran forest
pests in Bulgaria. Acta Zool. Bulg. 2013, 65, 179–186.
37. Sevim, A.; Demir, I.; Demirbaǧ, Z. Molecular Characterization and Virulence of Beauveria spp. from the Pine Processionary Moth,
Thaumetopoea pityocampa (Lepidoptera: Thaumetopoeidae). Mycopathologia 2010, 170, 269–277. https://doi.org/10.1007/s11046-
010-9321-6.
38. Pacheco, S.; Gómez, I.; Chiñas, M.; Sánchez, J.; Soberón, M.; Bravo, A. Whole Genome Sequencing Analysis of Bacillus thurin-
giensis GR007 Reveals Multiple Pesticidal Protein Genes. Front. Microbiol. 2021, 12, 758314.
https://doi.org/10.3389/fmicb.2021.758314.
39. Pang, A.S.D.; Gringorten, J.L.; Bai, C. Activation and fragmentation of Bacillus thuringiensis δ-endotoxin by high concentrations
of proteolytic enzymes. Can. J. Microbiol. 1999, 45, 816–825. https://doi.org/10.1139/w99-086.
40. Rukmini, V.; Reddy, C.Y.; Venkateswerlu, G. Bacillus thuringiensis crystal δ-endotoxin: Role of proteases in the conversion of
protoxin to toxin. Biochimie 2000, 82, 109–116. https://doi.org/10.1016/S0300-9084(00)00355-2.
41. Ivanov, K.; Stoimenova, A.; Obreshkova, D.; Saso, L. Biotechnology in the production of pharmaceutical industry ingredients:
Amino acids. Biotechnol. Biotechnol. Equip. 2013, 27, 3620–3626. https://doi.org/10.5504/bbeq.2012.0134.
42. Zhang, X.; Xu, G.; Shi, J.; Koffas, M.A.G.; Xu, Z. Microbial Production of L-Serine from Renewable Feedstocks. Trends Biotechnol.
2018, 36, 700–712. https://doi.org/10.1016/j.tibtech.2018.02.001.
43. Hess, S.; Van Beek, J.; Pannell, L.K. Acid hydrolysis of silk fibroins and determination of the enrichment of isotopically labeled
amino acids using precolumn derivatization and high-performance liquid chromatography-electrospray ionization-mass spec-
trometry. Anal. Biochem. 2002, 311, 19–26. https://doi.org/10.1016/S0003-2697(02)00402-5.
44. Fountoulakis, M.; Lahm, H.W. Hydrolysis and amino acid composition analysis of proteins. J. Chromatogr. A 1998, 826, 109–134.
https://doi.org/10.1016/S0021-9673(98)00721-3.
45. Bartolomeo, M.P.; Maisano, F. Validation of a reversed-phase HPLC method for quantitative amino acid analysis. J. Biomol. Tech.
2006, 17, 131–137.
46. Sun, S.W.; Lin, Y.C.; Weng, Y.M.; Chen, M.J. Efficiency improvements on ninhydrin method for amino acid quantification. J.
Food Compos. Anal. 2006, 19, 112–117. https://doi.org/10.1016/j.jfca.2005.04.006.
47. Hopwood, D.A.; Bibb, M.J.; Chater, K.F.; Bruton, C.J.; Kieser, H.M.; Lydiate, D.; Smith, C.P.; Ward, J.M., S.H. Genetic Manipula-
tion of Streptomyces: A Laboratory Manual.; The John Innes Institute: Norwich, UK, 1985.
48. Lane, D.J. 16S/23S rRNA Sequencing. In Nucleic Acid Techniques in Bacterial Systematics; Stackebrandt, E., Goodfellow, M., Eds.;
John Wiley and Sons: New York, NY, USA, 1991.
Int. J. Mol. Sci. 2022, 23, 15253 23 of 23
49. Kim, O.S.; Cho, Y.J.; Lee, K.; Yoon, S.H.; Kim, M.; Na, H.; Park, S.C.; Jeon, Y.S.; Lee, J.H.; Yi, H.; et al. Introducing EzTaxon-e: A
prokaryotic 16s rRNA gene sequence database with phylotypes that represent uncultured species. Int. J. Syst. Evol. Microbiol.
2012, 62, 716–721. https://doi.org/10.1099/ijs.0.038075-0.
50. Tamura, K.; Stecher, G.; Peterson, D.; Filipski, A.; Kumar, S. MEGA6: Molecular Evolutionary Genetics Analysis Version 6.0.
Mol. Biol. Evol. 2013, 30, 2725–2729. https://doi.org/10.1093/molbev/mst197.
51. Bolger, A.M.; Lohse, M.; Usadel, B. Trimmomatic: A flexible trimmer for Illumina sequence data. Bioinformatics 2014, 30, 2114–
2120. https://doi.org/10.1093/bioinformatics/btu170.
52. Prjibelski, A.; Antipov, D.; Meleshko, D.; Lapidus, A.; Korobeynikov, A. Using SPAdes De Novo Assembler. Curr. Protoc.
Bioinform. 2020, 70, e102. https://doi.org/10.1002/CPBI.102.
53. Gurevich, A.; Saveliev, V.; Vyahhi, N.; Tesler, G. QUAST: Quality assessment tool for genome assemblies. Bioinformatics 2013,
29, 1072–1075. https://doi.org/10.1093/bioinformatics/btt086.
54. Manni, M.; Berkeley, M.R.; Seppey, M.; Simão, F.A.; Zdobnov, E.M. BUSCO Update: Novel and Streamlined Workflows along
with Broader and Deeper Phylogenetic Coverage for Scoring of Eukaryotic, Prokaryotic, and Viral Genomes. Mol. Biol. Evol.
2021, 38, 4647–4654. https://doi.org/10.1093/molbev/msab199.
55. Hyatt, D.; Chen, G.L.; LoCascio, P.F.; Land, M.L.; Larimer, F.W.; Hauser, L.J. Prodigal: Prokaryotic gene recognition and trans-
lation initiation site identification. BMC Bioinform. 2010, 11, 119. https://doi.org/10.1186/1471-2105-11-119.
56. Camacho, C.; Coulouris, G.; Avagyan, V.; Ma, N.; Papadopoulos, J.; Bealer, K.; Madden, T.L. BLAST+: Architecture and appli-
cations. BMC Bioinform. 2009, 10, 421. https://doi.org/10.1186/1471-2105-10-421.
57. The Uniprot Consortium. UniProt: The universal protein knowledgebase in 2021. Nucleic Acids Res. 2021, 49, D480–D489.
https://doi.org/10.1093/nar/gkaa1100.
58. R Core Team. R: A Language and Environment for Statistical Computing; R Foundation for Statistical Computing: Vienna, Austria,
2022.
59. Carbon, S.; Ireland, A.; Mungall, C.J.; Shu, S.; Marshall, B.; Lewis, S.; Lomax, J.; Mungall, C.; Hitz, B.; Balakrishnan, R.; et al.
AmiGO: Online access to ontology and annotation data. Bioinformatics 2009, 25, 288–289. https://doi.org/10.1093/bioinformat-
ics/btn615.
60. Blum, M.; Chang, H.Y.; Chuguransky, S.; Grego, T.; Kandasaamy, S.; Mitchell, A.; Nuka, G.; Paysan-Lafosse, T.; Qureshi, M.;
Raj, S.; et al. The InterPro protein families and domains database: 20 years on. Nucleic Acids Res. 2021, 49, D344–D354.
https://doi.org/10.1093/nar/gkaa977.
61. Bertazzo, S. Electron Microscopy. In Microscopy of the Heart; Kaestner, L., Lipp, P., Eds.; Methods in Molecular Biology; Springer
International Publishing: Cham, Switzerland, 2018; Volume 1117, pp. 119–132; ISBN 9783319953045.
62. Holmes, D.S.; Quigley, M. A Rapid Boiling Method for the Preparation of Bacterial Plasmids. Anal. Biochem. 1981, 114, 193–197.
63. Green, R., M.; Sambrook, J. Molecular Cloning: A Laboratory Manual, 4th ed.; Cold Spring Harbor Laboratory Press: New York,
NY, USA, 2012; ISBN 9781936113415.
64. Jakob, F.; Lehmann, C.; Martinez, R.; Schwaneberg, U. Increasing protein production by directed vector backbone evolution.
AMB Express 2013, 3, 39. https://doi.org/10.1186/2191-0855-3-39.
65. Tautz, D.; Renz, M. An optimized freeze-squeeze method for the recovery of DNA fragments from agarose gels. Anal. Biochem.
1983, 132, 14–19. https://doi.org/10.1016/0003-2697(83)90419-0.
66. Xue, G.P.; Johnson, J.S.; Dalrymple, B.P. High osmolarity improves the electro-transformation efficiency of the gram-positive
bacteria Bacillus subtilis and Bacillus licheniformis. J. Microbiol. Methods 1999, 34, 183–191. https://doi.org/10.1016/S0167-
7012(98)00087-6.
67. Untergasser, A.; Cutcutache, I.; Koressaar, T.; Ye, J.; Faircloth, B.C.; Remm, M.; Rozen, S.G. Primer3-new capabilities and inter-
faces. Nucleic Acids Res. 2012, 40, e115. https://doi.org/10.1093/nar/gks596.