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Applied Physics A (2020) 126:368
https://doi.org/10.1007/s00339-020-03533-2
T.C. BIOLOGICAL ANDBIOMIMETIC MATERIALS
Electrochemical method forisolation ofchitinous 3D scaolds
fromcultivated Aplysina aerophoba marine demosponge andits
biomimetic application
KrzysztofNowacki1· IzabelaStępniak1· TomaszMachałowski2,3· MarcinWysokowski2,3· IaroslavPetrenko3·
ChristophSchimpf4· DavidRafaja4· EnricoLanger5· AndreasRichter5· JerzyZiętek6· SnežanaPantović7·
AlonaVoronkina8· ValentineKovalchuk9· ViatcheslavIvanenko10,11· YuliyaKhrunyk12,13· RobertaGalli14·
YvonneJoseph3· MichaelGelinsky15· TeolJesionowski2· HermannEhrlich3,16
Received: 16 March 2020 / Accepted: 7 April 2020
© The Author(s) 2020
Abstract
Three-dimensional (3D) biopolymer-based scaffolds including chitinous matrices have been widely used for tissue engi-
neering, regenerative medicine and other modern interdisciplinary fields including extreme biomimetics. In this study, we
introduce a novel, electrochemically assisted method for 3D chitin scaffolds isolation from the cultivated marine demosponge
Aplysina aerophoba which consists of three main steps: (1) decellularization, (2) decalcification and (3) main deproteiniza-
tion along with desilicification and depigmentation. For the first time, the obtained electrochemically isolated 3D chitinous
scaffolds have been further biomineralized exvivo using hemolymph of Cornu aspersum edible snail aimed to generate
calcium carbonates-based layered biomimetic scaffolds. The analysis of prior to, during and post-electrochemical isolation
samples as well as samples treated with molluscan hemolymph was conducted employing analytical techniques such as
SEM, XRD, ATR–FTIR and Raman spectroscopy. Finally, the use of described method for chitin isolation combined with
biomineralization exvivo resulted in the formation of crystalline (calcite) calcium carbonate-based deposits on the surface
of chitinous scaffolds, which could serve as promising biomaterials for the wide range of biomedical, environmental and
biomimetic applications.
Keywords Chitin· Scaffolds· Electrolysis· Biomineralization· Biomimetics· Hemolymph· Marine sponges· Aplysina
aerophoba
1 Introduction
In recent decades, the synthesis and application of 3D
biopolymer-based scaffolds represent one of the new trends
in environmental science and technology. Owing to their
ability to mimic the patterns of natural structures, excellent
biocompatibility, high biodegradability and non-toxicity 3D
scaffolds of natural origin find increasing applications in
medicine, biotechnology and various interdisciplinary fields
including tissue engineering, biomimetics, biocatalysis,
adsorption techniques and wastewater treatment [1–10].
Highly versatile and promising biopolymers such as cellu-
lose, chitin, collagen and their derivatives are more and more
frequently used in modern technology [11–16].
Chitin is one of the most abundant polysaccharides of
natural origin, obtained mainly from crustaceans, although
it can be found in representatives of diatoms, sponges,
mollusks, tubeworms, insects and arachnids [17–22].
This biopolymer is composed of β-(1,4)-N-acetyl-
d
-
glucosamine units and plays a crucial role in the forma-
tion of both soft and mineralized skeletal structures in
invertebrates requiring rigidity and mechanical strength
[23, 24]. Usually, chitin as a source of industrially pro-
duced chitosan is isolated by two main types of extraction
process: chemical and biological methods. The schematic
view showing principal steps of each method is repre-
sented in Fig.1 [25, 26]. In brief, industrial chemical
* Krzysztof Nowacki
krzysztof.j.nowacki@doctorate.put.poznan.pl
* Izabela Stępniak
izabela.stepniak@put.poznan.pl
Extended author information available on the last page of the article
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K.Nowacki et al.
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368 Page 2 of 16
treatment involves three major steps: deproteinization,
demineralization and depigmentation. Protein hydrolysis
occurring in the first step is carried out using alkali solu-
tions such as NaOH or KOH. In this stage of the isolation
process, the efficiency mostly depends on the concentra-
tion of a base, temperature and duration of treatment [27].
It should be also noted that for proper deproteinization of
chitin precursor a great excess of the alkali solution must
be used. The second step is usually completed by treat-
ing the source of chitin with acid solution (CH3COOH
or HCl) to ensure elimination of calcium carbonates. In
order to isolate chitinous skeleton, deproteinization and
demineralization steps are often repeated. The final treat-
ment step, depigmentation, is performed by adding highly
reactive oxidizing agents such as hydrogen peroxide. A
major drawback of the chemical isolation is caused by the
use of hazardous to the environmental extraction agents.
Moreover, these chemicals are used in great excess, gen-
erating effluents that must be neutralized prior to disposal
[28–30]. To prevent this negative environmental effect,
the biological/enzymatic treatment was developed as an
alternative method of chitin isolation [31–33]. Accord-
ing to this method, chemical extraction steps are substi-
tuted by the action of microorganisms and enzymes (see
Fig.1). Though requiring more time, such biological treat-
ment results in the isolation of chitin with a better pre-
served spatial structure [34]. Notwithstanding, in order to
increase the efficiency of biological method and reduce the
environmental impact of chemical process, the novel and
modified methods of chitin isolation have been developed
in recent time [35–38].
Marine demosponges have been recognized as a novel
source of naturally prefabricated 3D matrices and remain
to be the subject of intense research with respect to design-
ing of effective approaches to extract ready-to-use chitin-
ous scaffolds [38–40]. For this purpose, a well-established
chemical method was developed and since this treatment
involves acidic and alkali extraction steps that are cyclically
repeated, the process duration often exceeds 72h [41, 42].
Therefore, this method is often modified in order to reduce
treatment time and the amount of chemicals that are used.
Recently reported methods of chitin extraction are mostly
focused on the use of microwave irradiation as accelerating
factor [43, 44].
The only proposed approach including electrochemical
treatment of a chitin precursor was described previously by
Prof. Kuprina group [45–49]. The principle of this method is
based on the electrolysis of diluted NaCl aqueous solution to
ensure acidic and alkali treatment of crustacean’s Gammarus
pulex (Linnaeus, 1758) biomass [49].
Electrolysis is a well-known electrochemical process that
is thermodynamically forced by the flow of direct electric
current from an external source [50–52]. For this process
to function, a specific electrolytic cell (electrolyzer) must
be constructed: Briefly, the apparatus is composed from
two chambers and two symmetrical polarizable electrodes
Fig. 1 Schematic diagram
showing the isolation of chitin
(for details see Refs. [25, 26])
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made from chemically inert materials with high surface area.
Usually both electrodes are dipped in liquid electrolyte and
separated with an ion exchange membrane (cation, anion
or bipolar) [53–62]. Electrolysis of aqueous salts solutions
(NaCl or Na2SO4) in ion exchange membrane electrolyzers
is a relatively easy method to produce alkali and acids. One
of the most popular processes in industry is electrolysis of
Na2SO4 aqueous solution in a cation exchange membrane
(CEM) electrolyzer [60]. The basic principle of this process
is shown in Fig.2 [61].
In brief, the conversion of Na2SO4 to NaOH and H2SO4
can be separated in two steps: (1) electrochemical decompo-
sition of water particles and (2) separation of sodium cations
and sulfate anions [58]. The first step occurs on the anode
surface:
and on the cathode surface:
These redox reactions result in excess of H+ ions in elec-
trolyte solution in anode chamber (anolyte) and OH– ions in
cathode chamber (catholyte). Owing to this phenomenon,
boosted by synergic effect of the separation of sodium cati-
ons and sulfate anions on the CEM membrane, it is possible
to establish and change pH in each part of electrolyzer by
applying specific potential conditions.
(1)
H2
O→O
2
+H
+
+4e
−
(2)
4H2O
+
4e−
→
2H2
+
4OH−
Electrolysis has never been used for the isolation of
sponge chitin. In this study, for the first time, we applied con-
centrated Na2SO4 aqueous solution as electrolyte for a novel
electrochemical method which was designed by combining
well-known chemical treatment of chitin-based skeleton of
cultivated under marine farming conditions [63] Aplysina
aerophoba (Nardo 1833) marine demosponges with insitu
electrolysis. Finally, we used electrochemically isolated 3D
chitinous scaffolds for their biomimetic biomineralization
exvivo using hemolymph of industrially cultivated edible
snail Cornu aspersum with the aim of developing calcium
carbonate based scaffolds potentially applicable for environ-
mental remediation.
2 Materials andmethods
2.1 Biological samples andchemicals
2.1.1 Aplysina aerophoba demosponges
Selected specimens of cultivated A. aerophoba (Nardo,
1833) marine demosponges [63] in the form of air-dried
material (see Fig.3) were purchased from BromMarin
GmbH, Freiberg, Germany. Sodium sulfate [≥ 99.7% (VWR,
Dresden, Germany)] was used for the preparation of aqueous
electrolyte solution. All aqueous solutions were prepared
with distilled water.
Fig. 2 Schematic illustration of the electrolysis cell assembled in this study and a general principle of Na2SO4 aqueous solution electrolysis [61]
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K.Nowacki et al.
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368 Page 4 of 16
2.1.2 Cornu aspersum snails
One-year-old snails C. aspersum were obtained from
a commercial heliculture farm from the central part of
Poland (Snails Breeder—Hodowla Ślimaków, Małaszyce,
Poland). Snails were maintained in a special glass aquarium
at room temperature. As food, lettuce, carrots and apples
were traditionally used. Additionally, cuttlebone of Sepia
officinalis cuttlefish was given to snails as calcium carbonate
enrichment (CaCO3) in the diet. The snails were kept moist
throughout the experimental period with wet humus. Animal
rights statement is not required.
2.2 Non‑lethal hemolymph sampling procedure
For hemolymph collection, the modified non-lethal intravital
method described previously [64, 65] was used. The shell
surface of the selected C. aspersum snail was cleaned by
70% ethanol. After that, a piece of shell was removed and
3h later about 0.5ml of hemolymph was isolated from an
individual of C. aspersum by main vessel puncture using
a sterile 1ml syringe and needle of 0.45mm in diameter
[64, 65] (see Fig.4). Twelve hours post-isolation, an organic
film was observed in the site of the removed shell fragment,
which was fully mineralized within the next few days. Dur-
ing the next four months after the procedure, the snails used
in this study did not show any visible changes neither in their
physiology nor in behavior.
2.3 Electrolytic cell setup
Schematic diagram of the experimental setup for the electro-
chemically assisted isolation of chitin is depicted in Fig.2.
The CEM (cation exchange membrane) electrolyzer con-
sisted of two cylindrical poly(propylene) chambers (50ml
each) separated by a cellulose membrane made from filter
paper (75gcm−2 (ChemLand, Stargard, Poland)) and sealed
with parafilm (Bemis Company Inc., Neenah, USA). Elec-
trodes (cathode and anode) were made of platinum sheets
(effective area: 2.2 cm2). Distance between both electrodes
was about 10.0cm and they were connected with DC power
supply [VoltCraft PS2043D (Conrad Electronic Group,
Hirschau, Germany)] by platinum wire current collectors.
Fig. 3 Fragment of A. aerophoba demosponge from the marine
ranching facility in Kotor Bay, Montenegro (a, b). Mineralized and
pigmented skeletal fibers (c) remain to be rigid after drying on air at
28°C
Fig. 4 Non-lethal isolation of the hemolymph from C. aspersum
snail. Initially, a small fragment of the shell (around 5mm × 5mm)
was carefully removed using scalpel with the aim to obtain an access
to the main vessel (a). Common view of the procedure of non-lethal
hemolymph aspiration using sterile syringe and needle (b)
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1.9M sodium sulfate aqueous solution with an initial tem-
perature of 40°C was utilized as anolyte as well as catholyte.
2.4 Electrochemically assisted isolation ofsponge
chitin
The isolation of chitin scaffold from A. aerophoba was per-
formed by novel electrochemically assisted method (see
Fig.5) proposed in this study. In the pretreatment step, 0.2g
A. aerophoba sample was rinsed repeatedly with distilled
water for 24h (25°C) in order to remove water-soluble salts
and various major impurities. Water-swelled sample, free
from impurities and with partially lysed cells, was ready
for treatment in electrolyzer. It should to be noted that after
each step the sample was extensively washed and stirred in
distilled water till neutral pH. Also, the initial concentration
of electrolyte (Na2SO4) for every step was 1.9moll−1 and
start temperature for both anolyte and catholyte was 40°C.
Electrochemical treatment was performed in three main
steps (see Fig.6).
Step 1 Decellularization (predeproteinization) was car-
ried out in the cathode chamber for 3h (12V; 0.5 A;
50°C). High pH of the catholyte (up to 12.0) caused a
complete lysis of preswelled cells and a partial degrada-
tion of lipids and proteins which resulted in the removal
of soft tissues. Post-treated sample was composed of deep
brown cell-free skeleton.
Step 2 Decalcification (demineralization) was per-
formed in the anode chamber for 3h (12V; 0.7 A; 50°C).
Low pH (down to 1.5) along with free access of the anolyte
solution to the sponge skeleton resulted in the dissolution
of calcium and magnesium carbonate, acid-soluble pig-
ments and proteins. Post-treated sample was in form of a
light yellow cell-free skeleton.
Step 3 Main deproteinization, desilicification and depig-
mentation were conducted in the cathode chamber for 3h
(16V; 1.5 A; 60°C). Extremely high pH (up to 12.5) of
the catholyte caused by increased electrolysis current was
used to completely remove pigments and residual proteins
from the chitinous matrix. Post-treated sample presented
a colorless scaffold. After treatment in the electrolyzer,
the sample was once more extensively rinsed with dis-
tilled water until neutral pH and stored in ethanol absolute
(4°C).
Fig. 5 Schematic view of the electrochemically assisted isolation of
chitin scaffold from A. aerophoba
Fig. 6 Experimental setup for all steps of the electrochemically
assisted isolation of sponge chitin in the form of 3D scaffold
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K.Nowacki et al.
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2.5 Biomineralization exvivo ofelectrochemically
isolated A. aerophoba demosponge chitinous
scaolds
A square fragment (10 × 10mm) of selected chitinous scaf-
fold was immersed in hemolymph of C. aspersum for one
hour and then placed on a sterile slide until completely dried
at room temperature. This procedure was repeated for five
times. The described technique aimed to simulate biominer-
alization of organic matrix by living snail under natural con-
ditions via mimicking of physicochemical effects reported
for shell regeneration of terrestrial snails previously [66–68].
Obtained in this way, mineral phases deposited on chitinous
matrix were characterized as represented below.
2.6 Short‑term cultivation ofhemocytes
onchitinous matrix
For short-term cultivation of hemocytes about 0.5ml of
the C. aspersum hemolymph was isolated using the method
described above and placed in 2-ml Eppendorf vessel.
Selected fragment of electrochemically isolated chitinous
matrix as represented in Fig.7n was immersed in the hemo-
lymph at room temperature for 24h. To prevent possible
bacterial contamination, streptomycin [100μgml−1 (Merck,
Darmstadt, Germany)] and penicillin [60μgml−1 (Merck,
Darmstadt, Germany)] were used [69].
2.7 Characterization ofobtained materials
2.7.1 Photography andgures
Photographs and macroscopic images were performed by
Nikon D-7100 camera with Nikon AF-S DX 18–105mm
f/3.5–5.6G and Nikon AF-S VR Micro-Nikkor 105mm
f/2.8G IF-ED objective lenses. Figures were prepared using
the GNU Image Manipulation Program GIMP and the
Microsoft Office tool PowerPoint 2016.
2.7.2 Digital, light anduorescence microscopy
The samples were observed using advanced imaging and
measurement system consisting of Keyence VHX-6000 digi-
tal optical microscope and the swing-head zoom lenses VH-
Z20R (magnification up to 200×) and VH-Z100UR (magni-
fication up to 1000×) (Keyence, Osaka, Japan). The light and
fluorescence microscopy mode was performed by Keyence
BZ-9000 (Keyence, Osaka, Japan) microscope. Calcite min-
eral standard has been purchased from International Institute
of Biomineralogy (INTIB GmbH, Freiberg, Germany).
2.7.3 Eosin andmethylene blue staining
Hemavet (Kolchem, Łódź, Poland), the combination of eosin
and methylene blue dyes, was used. Previously, this stain
was successfully used for hemocyte detection and characteri-
zation [70, 71] Hemolymph cell monolayers (HCMs) were
prepared by spreading the drop of hemolymph on sterile
glass slide and drying it at ambient temperature. Staining
was also used to detect hemocytes settled on chitinous scaf-
fold (see Fig.9).
2.7.4 Alizarin Red S staining
Ex vivo mineralized chitinous scaffolds were stained with
Alizarin Red S (Sigma-Aldrich, Taufkirchen, Germany) and
compared with that of native A. aerophoba chitin as iso-
lated. For the staining procedure 40mM of Alizarin Red S
(pH 8.3) was used for staining of the samples during 30min
at room temperature (for details see [72]). Stained samples
were washed with distilled water for ten times to eliminate
the unattached Alizarin Red S as well as mineral particles
which were not tightly attached to the surface of chitin fib-
ers. Calcium deposits were detected as orange–red color
microagglomerates employing digital microscopy.
2.7.5 ATR–FTIR spectroscopy
Infrared spectroscopy techniques were used for the qualita-
tive characterization of obtained mineralized scaffolds as
well as pure chitin isolated from A. aerophoba. The presence
of expected functional group was confirmed by ATR–FTIR
(attenuated total reflectance–Fourier transform infrared
spectroscopy) and verified using Nicolet 210c spectrometer
(Thermo Scientific, Waltham, USA). The investigation was
performed over a wave number range of 1900–500cm−1
(resolution of 0.5cm−1).
2.7.6 Raman spectroscopy
Raman spectra were recorded using a Raman spectrometer
RamanRxn1 (Kaiser Optical Systems Inc., Ann Arbor, USA)
coupled to a light microscope DM2500 P (Leica Microsys-
tems GmbH, Wetzlar, Germany). The excitation of Raman
scattering was obtained with a diode laser emitting at a
wavelength of 785nm, propagated to the microscope with a
100µm optical fiber and focused on the samples by means
of a 50×/0.75 microscope objective, leading to a focal spot
of about 20µm with a power of 170 mW. The Raman sig-
nal was collected in reflection configuration and sent to the
f/1.8 holographic imaging spectrograph by using 62.5µm
core optical fiber. The spectral resolution in the range of
150–3250cm−1 was 4cm−1. Raman spectra were punctu-
ally recorded, using integration time of 1s and averaging
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Electrochemical method forisolation ofchitinous 3D scaffolds fromcultivated Aplysina…
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60 spectra in order to improve the signal-to-noise ratio. A
variable baseline was calculated to remove the background
by applying the function “msbackadj” of theMATLAB tool-
boxes(MathWorks Inc., Natick, USA). This baseline was
estimated within multiple windows of 150cm−1 width and
shifted with 150cm−1 step and using a linear interpolation
method.
Raman spectra were recorded using a Raman spectrom-
eter RamanRxn1 (Kaiser Optical Systems Inc., Ann Arbor,
USA) coupled to a light microscope DM2500 P (Leica
Microsystems GmbH, Wetzlar, Germany). The excitation of
Raman scattering was obtained with a diode laser emitting at
a wavelength of 785nm, propagated to the microscope with
a 100µm optical fiber and focused on the samples by means
of a 50×/0.75 microscope objective, leading to a focal spot
of about 20µm with a power of 170 mW. The Raman sig-
nal was collected in reflection configuration and sent to the
f/1.8 holographic imaging spectrograph by using 62.5µm
core optical fiber. The spectral resolution in the range of
150–3250cm−1 was 4cm−1. Raman spectra were punctually
Fig. 7 A. aerophoba sample
prior to (a–c) and at different
stages of electrochemically
assisted isolation of chitinous
3D scaffolds: 1.5h (d–g) and
3h (h–j) starting from the first
catholyte treatment; 3h fol-
lowing anolyte treatment (k, l)
and 3h post-second catholyte
treatment (m–o)
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K.Nowacki et al.
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368 Page 8 of 16
recorded, using integration time of 1s and averaging 60
spectra in order to improve the signal-to-noise ratio. A vari-
able baseline was calculated to remove the background by
applying the function “msbackadj” of theMATLAB tool-
boxes(MathWorks Inc., Natick, USA). This baseline was
estimated within multiple windows of 150cm−1 width and
shifted with 150cm−1 step and using a linear interpolation
method.
2.7.7 X‑ray diraction (XRD)
The phase composition of samples was analyzed by X-ray
diffraction using SEIFERT-FPM URD6 diffractometer
equipped with a sealed X-ray tube with Cu anode and a
secondary graphite monochromator placed in front of a
proportional counter. For the measurement, the sample was
fixed on its sides with scotch tape to a “zero background”
to a sample holder, assuming that the amount of tape in the
beam was small. The phase identification was performed
with the ICDD PDF-4 + database linked to PANalytical
HighScore + software. The Rietveld refinement [73] for a
more detailed analysis of the sample was conducted employ-
ing Maud software package [74].
2.7.8 Scanning electron microscopy (SEM)
The specimens were fixed on an aluminum sample holder
with conductive carbon adhesive tabs and were sputtered
with platinum for 15s at a distance of 30mm by an Edwards
S150B sputter coater. The scanning electron micrographs
were observed using a high-resolution Hitachi S-4700-II
(Hitachi, Ltd., Tokyo, Japan) equipped with a cold field
emission gun. The elements were analyzed by energy-dis-
persive X-ray spectroscopy in the EDX analysis system from
EDAX and XL30ESEM Philips—scanning electron micro-
scope (Philips, Amsterdam, the Netherlands).
3 Results anddiscussion
The main goal of this study was to develop a fast, low-cost
and low-effluent method of 3D chitin isolation, based on in
situ electrolysis of aqueous Na2SO4 solution. The concept
of electrochemically assisted isolation process was split into
three main steps. Morphological changes within A. aero-
phoba sample were analyzed by digital optical microscope
and SEM. Figure7a–c shows a small cutoff of the examined
sample prior to treatment in the electrolytic cell. The pho-
tographs of the sample taken right after pretreatment step
(washing in distilled water) demonstrated a well-preserved
original bio-architecture of A. aerophoba sponge (tissue
structures supported by chitinous scaffold). Next, in order
to decellularize the sample the electrolytic treatment was
applied (see Figs.5, 6). Indeed, the alkaline environment of
the catholyte solution should have resulted in the complete
lysis of A. aerophoba cells. Photographic images of this pro-
cess taken after 1.5 h since the beginning of the first electro-
chemically assisted isolation step (12 V; 0.5 A) are presented
in Fig.7d–g. The microscopic investigation of this sample
revealed that cells and tissues had decomposed during catho-
lyte treatment. Moreover, the dissolution of A. aerophoba
somatic cells proceeded gradually from the external area
of the sample to internal chitinous scaffold which is clearly
visible in Fig.7e, g. The sample cutoff after full-time catho-
lyte treatment (3 h; 12 V; 0.5 A) is depicted in Fig.7h–j.
This image illustrates dark-brown, semitransparent and cell-
free chitinous scaffold, the spatial structure of which can
be characterized as typical branched network of chitinous
tubes of the sponge origin. In a higher magnification, a few
residues of tissue fragments can be still observed (Fig.7j);
however, the time needed to achieve this level of decompo-
sition effect was incomparably shorter in comparison with
previously reported standard methods [1, 3]. The second
step of the electrochemically assisted isolation method was
applied in order to remove possible carbonate salts within
the sample. However, since A. aerophoba sponge minerali-
zation was insignificant, this stage of the process was carried
out mostly to remove acid-soluble pigments and proteins.
The cutoff of the sample after full-time anolyte treatment
(3 h; 12 V; 0.7 A) is presented in Fig.7k, l. The treated
sample after the second electrochemically assisted isolation
step appeared in the form of a light yellow, cell-free 3D
skeleton with a well preserved spatial structure consisting
of branched microtubular network. The final treatment was
performed to get rid of the remained proteins, pigments and
possible silica remnants. Due to structural incorporation of
these compounds into the chitinous tubes [21, 24], the third
electrochemically assisted isolation step was conducted in
extremely corrosive conditions (3 h; 16 V; 1.5 A; 60 °C).
Figure7m-o shows the sample after full electrolytic treat-
ment presented the form of a colorless scaffold with slightly
harmed spatial structure of the branched tubular network.
The partial damage of the spatial structure of chitinous tubes
was probably caused by intensive gas evolution on the cath-
ode. The black areas in Fig.7m are artificial and became
visible through the focus on the fibers surface. Additionally,
we conducted SEM analysis (Fig. 8) of the electrochemically
treated A. aerophoba skeleton fragments similar to those
represented in Fig.7. Corresponding EDX analysis (Fig.8c,
d) was carried out to monitor changes within the chemical
composition of treated matter with respect to diverse ele-
ments usually belonging to skeletal fibers of A. aerophoba
[21]. The nature of obtained scaffold was characterized using
FTIR–ATR and XRD (see Figs.12, 14, respectively). The
identification of chitin confirmed that intensive deacetyla-
tion process and transformation of chitin into chitosan had
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not occurred under experimental conditions even after the
treatment in harsh catholyte environment.
Previously, such kinds of 3D chitinous scaffolds isolated
from diverse demosponges had been used in tissue engineer-
ing [1, 3, 75, 76] and extreme biomimetics [40, 77] However,
in this study we took the decision to mineralize the electro-
chemically isolated scaffolds of A. aerophoba origin using
mollusk hemolymph as a unique biological system which is
responsible for calcification of mechanically damaged shells
invivo.
It is well recognized that molluscan hemolymph con-
tains calcium Ca2+, bicarbonate HCO3− and other ions
(i.e. Na+, K+, Cl−) [78]. Apart from these ions, molluscan
hemolymph is also rich in cells and enzymes involved in
CaCO3 formation such as carbonic anhydrase [79–81].
Hemocytes, primarily involved in immunoreactions, are
also responsible for calcium-rich granules deposition dur-
ing the formation as well as regeneration of mollusks shell
[82, 83]. Eosin + methylene blue (Hemovet) is a standard
stain, primarily used for the differential staining of cellular
elements of blood. This protocol was used here to visual-
ize distribution of hematocytes on the chitin scaffold [71].
Our observations made with use of eosin + methylene blue
stain (see Fig.9) clearly indicate that hemocytes react with
chitin after 24h of immersion in the hemolymph from C.
aspersum snail. The formation of hemocyte-containing
clusters (Fig.9c) may be explained by the recognition of
chitin as a foreign body and nodulation exvivo. Moreover,
because chitin along with proteins form organic membrane
during shell regeneration [84–86] hemocytes may recog-
nize this amino polysaccharide as a natural scaffold useful
for biomineralization invivo with respect to the generation
of calcium carbonates. Such chitin-containing membrane
acts as a specialized scaffold for the granulocytes losing
their cytoplasm full of free nuclei and organic matter and
precipitating calcium-rich granules [82].
Alizarin Red S, an anthraquinone derivative, that may
be used to identify calcium in tissue sections. The reaction
is not strictly specific for calcium, since magnesium, man-
ganese, barium, strontium and iron may interfere, but these
elements usually do not occur in sufficient concentration
to interfere with the staining. Calcium forms an Alizarin
Red S–calcium complex in a chelation process, and the
end product is a bright red stain. Therefore, formation of
calcium-based crystals deposits on the surface of chitinous
scaffold after its immersion into C. aspersum hemolymph
could be easily confirmed by Alizarin Red S staining (see
Fig.10). Calcium deposits were detected using digital
microscopy in the form of red color agglomerates tightly
attached to the surface of isolated chitinous matrix. Hav-
ing assumed that snail uses physical- and cellular-based
processes [87–90] in shell regeneration, we aimed to ana-
lyze the polymorphs of the crystals structures obtained by
biomineralization exvivo.
Fig. 8 SEM imagery of the selected fragment of A. aerophoba
sponge skeleton prior to electrolytic treatment (a) after 1.5h of the
first catholyte treatment (b) and 3 h of the second catholyte treat-
ment (c) confirms structural changes represented in Fig. 7. EDX
analysis shows changes in the chemical composition of naturally
occurring skeletal fibers (d) [corresponding to SEM image (a)] and
electrochemically isolated chitinous scaffold (e) [corresponding to
SEM image (c)]. Residual amounts of Na and S (e) which originated
from Na2SO4 were finally removed from chitin using dialysis against
deionized water during 12h at room temperature
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K.Nowacki et al.
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368 Page 10 of 16
Additional analysis of the obtained deposits carried out by
fluorescence microscopy indicated similarity between cal-
cite standard [92, 93] and microgranular phase formed after
biomineralization exvivo using C. aspersum hemolymph
with respect to the red auto-fluorescence (see Fig.11). More
advanced techniques such as ATR–FTIR (Fig.12), Raman
spectroscopy (Fig.13) as well as X-ray diffraction analysis
(Fig.14) were further applied to identify calcium carbonate
polymorph formed.
Spectra obtained using ATR–FTIR analysis of the chitin-
ous scaffold before and after biomineralization exvivo are
demonstrated in Fig.12. Both spectra show a characteristic
band for α-chitin, such as amide I at 1633cm−1 (see gold and
grey lines). This band corresponds to the presence of stretch-
ing vibrations from intermolecular (C=O⋯HN) and intra-
molecular (C=O⋯HO(C6); C=O⋯HN) hydrogen bonds
[22, 94]. The presence of such bands as amide II (νN–H and
νC–N) at 1548–1538cm−1, amide III (νC–N and δN–H)
at 1308cm−1 or characteristic intense bands at 899cm−1
(C–O–C bridge as well as glycosidic linkage) additionally
proved the occurrence of α-chitin in the analyzed sample. A
sharp band visible at 873cm−1 (see blue arrow) undeniably
indicated that calcium carbonate (CaCO3) as monohydrocal-
cite polymorph [95–98] was formed at the chitinous scaffold
after biomineralization.
Raman spectra acquired from the obtained crystals con-
tain five bands (Fig.13). Four of the five fundamental modes
of calcite are visible at 279, 710, 1083 and 1433cm−1.
The first fundamental mode, normally observed at about
155cm−1, lies outside the acquired spectral range. The small
band at 1746cm−1, which is above the highest frequency of
the fundamental modes, should be attributed to an overtone
of the IR active mode observed at 873cm−1 [99].
The results obtained using ATR–FTIR were further con-
firmed by XRD, being the method of choice to analyze the
presence of certain crystal structures. The analysis of powder
X-ray diffraction patterns of the chitinous scaffold revealed
the presence of CaCO3 (calcite). Figure14 shows a SEM
micrograph of the calcium carbonate crystals, and the pat-
tern, displayed in Fig.14, compares the measured data with
the Rietveld-like refinement of the calcite phase together
with manually set background. The contributions of calcite
Fig. 9 Light microscopy imagery. Single (a) and aggregated (b)
hemocytes present in isolated hemolymph of C. aspersum snail can
be observed without (a) and using eosin and methylene blue staining
(b) on the glass slide. Visible pseudopodia (yellow arrows) and navy
blue color after staining indicate the presence of granulocytes (b).
The formation of hemocytes-based clusters on the surface of A. aer-
ophoba chitinous scaffold, after 24h immersion in the hemolymph,
prior (c) and after staining with eosin and methylene blue stain (d)
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Electrochemical method forisolation ofchitinous 3D scaffolds fromcultivated Aplysina…
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Page 11 of 16 368
to the diffraction pattern are indicated at the bottom of the
plot (Fig.14) as a proof of the positive identification. The
obtained lattice parameters of calcite, a=(4.973±0.002)Å
and c=(16.987±0.008)Å, are smaller than the tabulated
values for ambient conditions (a=4.987Å, c=17.058Å)
[ICDD PDF #04-012-0489]. However, some maxima in
the pattern could not be identified unambiguously. A large
hump at 2θ≈20° most likely corresponds to the α-chitin
110 reflection [4] since chitin belongs to the scaffold as was
also identified by ATR–FTIR. A second contribution to
Fig. 10 3D chitinous scaffolds
electrochemically isolated from
A. aerophoba prior to biominer-
alization exvivo using C. asper-
sum hemolymph are represented
in digital microscopy image (a).
Alizarin Red S staining of this
chitin scaffold resulted in the
appearance of slightly violet
color (b). The formation of
granular transparent calcium-
based deposits on the surface
of chitin after biomineralization
exvivo is well visible using
digital light microscopy (c).
These deposits became well vis-
ible (arrows) after Alizarin Red
S staining due to intensive red
coloration (d) which is indica-
tive of Ca-based structures [91]
Fig. 11 Both the mineral
constituents, obtained after
biomineralization exvivo of 3D
chitin scaffold (a) using hemo-
lymph of C. aspersum snail as
well as calcite mineral standard
(c), represent a high similarity
with respect to their strong red
auto- fluorescence (b and d,
respectively)
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K.Nowacki et al.
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368 Page 12 of 16
this maximum can originate from amorphous components
in the samples. Other maxima at 2θ≈27° and 2θ≈33°
must, unfortunately, remain unassigned here. The former
of the two might also be explained by a-chitin (102 maxi-
mum), if a preferred orientation of chitin in the scaffold is
assumed. However, no in-depth analysis of this condition
was undertaken.
Nowadays, data concerning practical application of
calcite as well as other calcium carbonate-based phases
in biomedicine and remediation in diverse environments
are well represented in the literature. For example, cal-
cium carbonate-based scaffolds have found application in
modern biomedicine as constructs, which improve osteoin-
ductive potential [100–102]. Intriguingly, natural calcium
carbonate layer created by molluscs invivo may serve as a
biocompatible interface between selected metals or alloys
used in medical devices or as implants in human body
[103].
Heavy metals present in water resources pose a high risk
of hazard for human health as well as the environment. It
is well recognized that calcite has a great ability to adsorb
heavy metal ions [104–108]. Indeed, natural calcite as lime-
stone [109] or in the form of waste chicken eggshells [110]
has been used for heavy metal removal. In comparison with
other calcium carbonate polymorphs, calcite represents the
highest affinity to lead (Pb2+) [111] and cadmium (Cd2+)
[112] ions. Also, a high ability of calcite for the remedia-
tion of arsenic (As) contamination from water solutions was
reported [105, 106, 113]. Furthermore, in 2011, Fukushi
and coworkers revealed that monohydrocalcite possesses a
significantly higher arsenic sorption capacity in comparison
with calcite [104]. Finally, the use of calcite as a substrate
for the removal of up to 2300 mg/l fluoride from contami-
nated groundwater was described by Turner and coworkers
[114].
Fig. 12 ATR–FTIR spectra of A. aerophoba chitin scaffold before
(gold line) and after biomineralization ex vivo (grey line) in the
region of 1900–500cm−1
Fig. 13 Representative graph depicting Raman spectrum of the min-
eral phase obtained on the surface of 3D chitinous scaffold after
biomineralization exvivo mediated by C. aspersum snail hemolymph
Fig. 14 SEM micrograph of the calcium carbonate crystals (a)
formed during the ex vivo biomineralization of chitinous scaffolds
using hemolymph of C. aspersum snail Powder diffraction pattern of
the chitinous scaffold after biomineralization exvivo (b). Measured
data are plotted by open dots, and the calculated intensity is high-
lighted by a solid line. The contribution of calcite (CaCO3) is shown
at the bottom clearly verifying the presence of this phase in the sam-
ple. Two unassigned peaks remain in the data (2θ ≈ 27°, 2θ ≈ 33°).
The maximum at 2θ ≈ 20°, included in the background, belongs to
the α-chitin scaffold (α-chitin 110 reflection) or an amorphous com-
ponent in the sample
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Electrochemical method forisolation ofchitinous 3D scaffolds fromcultivated Aplysina…
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Page 13 of 16 368
Herein, for the first time we performed electrochemical
isolation of natural 3D chitinous scaffolds from cultivated
A. aerophoba marine demosponge and biomineralized
them biomimetically ex vivo using mulluscan hemolymph
as a source of hemocytes and calcium. This unique method
developed by us allowed the generation of crystalline (cal-
cite) calcium carbonate-based layers, which could be useful
for both biomedical and environmental applications in the
future.
4 Conclusions
In the present work, for the first time the in situ electroly-
sis of 1.9 M Na2SO4 aqueous solution in CEM membrane
electrolyzer was utilized as isolation method of chitinous
scaffolds from A. aerophoba demosponge. The final result
of electrochemically assisted isolation of chitin was a color-
less scaffold. The digital light microscopic investigation of
this final product revealed that despite a possible mechanical
damage the general spatial structure of the sample preserved
its original interconnected network of unique microtubu-
lar nature. Further characterization of the isolated sample
with FTIR and EDX techniques proved that a pure chitin-
ous scaffold can be obtained by the application of described
method. The perspective of the biomineralization ex vivo
to be used in biomimetic fields which is represented in our
study includes diverse open questions concerning, for exam-
ple, the role of hemocytes in the generation of fine-tuned
microenvironment necessary for biocalcification ex vivo.
Without doubt, further studies on the mechanical proper-
ties of developed mineralized scaffolds aimed at practical
application in environmental remediation should be carried
out in the near future.
Acknowledgements This work was performed with the financial
support of Poznan University of Technology, Poland (Grant No.
0911/SBAD/0380/2019), as well as by the Ministry of Science and
Higher Education (Poland) as financial subsidy to PUT No. 03/32/
SBAD/0906. Krzysztof Nowacki was supported by the Erasmus Plus
program (2019). Also, this study was partially supported by the DFG
Project HE 394/3 and SMWK Project No. 02010311 (Germany).
Marcin Wysokowski is financially supported by the Polish National
Agency for Academic Exchange (PPN/BEK/2018/1/00071). Tomasz
Machałowski is supported by DAAD (Personal Ref. No. 91734605).
Yuliya Khrunyk is supported by the Russian Science Foundation (Grant
No. 18-13-00220).
Compliance with ethical standards
Conflict of interest Authors declare no conflict of interest.
Consent to participate All the coworkers have agreed to participate.
Consent for publication All the coworkers have agreed with the pub-
lication.
Open Access This article is licensed under a Creative Commons Attri-
bution 4.0 International License, which permits use, sharing, adapta-
tion, distribution and reproduction in any medium or format, as long
as you give appropriate credit to the original author(s) and the source,
provide a link to the Creative Commons licence, and indicate if changes
were made. The images or other third party material in this article are
included in the article’s Creative Commons licence, unless indicated
otherwise in a credit line to the material. If material is not included in
the article’s Creative Commons licence and your intended use is not
permitted by statutory regulation or exceeds the permitted use, you will
need to obtain permission directly from the copyright holder. To view a
copy of this licence, visit http://creat iveco mmons .org/licen ses/by/4.0/.
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Aliations
KrzysztofNowacki1· IzabelaStępniak1· TomaszMachałowski2,3· MarcinWysokowski2,3· IaroslavPetrenko3·
ChristophSchimpf4· DavidRafaja4· EnricoLanger5· AndreasRichter5· JerzyZiętek6· SnežanaPantović7·
AlonaVoronkina8· ValentineKovalchuk9· ViatcheslavIvanenko10,11· YuliyaKhrunyk12,13· RobertaGalli14·
YvonneJoseph3· MichaelGelinsky15· TeolJesionowski2· HermannEhrlich3,16
1 Institute ofChemistry andTechnical Electrochemistry,
Poznan University ofTechnology, ul. Berdychowo 4,
60-965Poznan, Poland
2 Institute ofChemical Technology andEngineering, Faculty
ofChemical Technology, Poznan University ofTechnology,
Berdychowo 4, 60965Poznan, Poland
3 Institute ofElectronics andSensor Materials, TU
Bergakademie Freiberg, Gustav-Zeuner Str. 3,
09599Freiberg, Germany
4 Institute ofMaterials Science, TU Bergakademie Freiberg,
09599Freiberg, Germany
5 Institute ofSemiconductors andMicrosystems, TU Dresden,
01062Dresden, Germany
6 Department ofEpizootiology andClinic ofInfectious
Diseases, Faculty ofVeterinary Medicine, University ofLife
Sciences, Głęboka 30, 20612Lublin, Poland
7 Faculty ofMedicine, University ofMontenegro, Kruševac
bb, 81000Podgorica, Montenegro
8 Department ofPharmacy, National Pirogov Memorial
Medical University, Vinnytsia21018, Ukraine
9 Department ofMicrobiology, National Pirogov Memorial
Medical University, Vinnytsya, Vinnytsia21018, Ukraine
10 Department ofInvertebrate Zoology, Biological Faculty,
Lomonosov Moscow State University, Moscow,
Russia119992
11 Taxonomy andSystematics Group, Naturalis Biodiversity
Center, 2300RALeiden, TheNetherlands
12 Ural Federal University, Mira Str. 19, Ekaterinburg,
Russia620002
13 The Institute ofHigh Temperature Electrochemistry,
Ural Branch oftheRussian Academy ofSciences,
Akademicheskaya Str. 20, Ekaterinburg, Russia620990
14 Clinical Sensoring andMonitoring, Department
ofAnesthesiology andIntensive Care Medicine, Faculty
ofMedicine, TU Dresden, 01307Dresden, Germany
15 Centre forTranslational Bone, Joint, Soft Tissue Research,
Medical Faculty andUniversity Centre forOrthopaedics
andTrauma Surgery, University Hospital Carl Gustav Carus
atTechnische Universität Dresden, 01307Dresden, Germany
16 Center forAdvanced Technology, Adam Mickiewicz
University, 61614Poznan, Poland
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