Automated freeze-thaw cycles for decellularization of tendon tissue - a pilot study
Roth et al. BMC Biotechnology
Automated freeze-thaw cycles for decellularization of tendon tissue - a pilot study
Susanne Pauline Roth 0 1
Sina Marie Glauche 0
Amelie Plenge 2
Ina Erbe 1
Sandra Heller 4
Janina Burk 0 1 3
0 Saxonian Incubator for Clinical Translation, University of Leipzig , Philipp-Rosenthal-Strasse 55, Leipzig 04103 , Germany
1 Large Animal Clinic for Surgery, University of Leipzig , An den Tierkliniken 21, Leipzig 04103 , Germany
2 Tierklinik Kaufungen , Pfingstweide 2, Kaufungen 34260 , Germany
3 Institute of Veterinary Physiology, University of Leipzig , An den Tierkliniken 7, Leipzig 04103 , Germany
4 Department of Pathology and Laboratory Medicine, Tulane University , New Orleans , USA
Background: Decellularization of tendon tissue plays a pivotal role in current tissue engineering approaches for in vitro research as well as for translation of graft-based tendon restoration into clinics. Automation of essential decellularization steps like freeze-thawing is crucial for the development of more standardized decellularization protocols and commercial graft production under good manufacturing practice (GMP) conditions in the future. Methods: In this study, a liquid nitrogen-based controlled rate freezer was utilized for automation of repeated freeze-thawing for decellularization of equine superficial digital flexor tendons. Additional tendon specimens underwent manually performed freeze-thaw cycles based on an established procedure. Tendon decellularization was completed by using non-ionic detergent treatment (Triton X-100). Effectiveness of decellularization was assessed by residual nuclei count and calculation of DNA content. Cytocompatibility was evaluated by culturing allogeneic adipose tissue-derived mesenchymal stromal cells on the tendon scaffolds. Results: There were no significant differences in decellularization effectiveness between samples decellularized by the automated freeze-thaw procedure and samples that underwent manual freeze-thaw cycles. Further, we inferred no significant differences in the effectiveness of decellularization between two different cooling and heating rates applied in the automated freeze-thaw process. Both the automated protocols and the manually performed protocol resulted in roughly 2% residual nuclei and 13% residual DNA content. Successful cell culture was achieved with samples decellularized by automated freeze-thawing as well as with tendon samples decellularized by manually performed freeze-thaw cycles. Conclusions: Automated freeze-thaw cycles performed by using a liquid nitrogen-based controlled rate freezer were as effective as previously described manual freeze-thaw procedures for decellularization of equine superficial digital flexor tendons. The automation of this key procedure in decellularization of large tendon samples is an important step towards the processing of large sample quantities under standardized conditions. Furthermore, with a view to the production of commercially available tendon graft-based materials for application in human and veterinary medicine, the automation of key procedural steps is highly required to develop manufacturing processes under GMP conditions.
Regenerative medicine; Tissue engineering; Tendon; Horse; Automation; Decellularization; Controlled rate freezer
Decellularization of natural tissues offers promising
opportunities as a multi-purpose tool in the field of tissue
engineering . Particularly, the challenging creation of
decellularized biological scaffolds with a preserved
tissuespecific extracellular matrix (ECM) composition provides
a crucial field of application. The importance of biological
scaffolds with a naturally structured ECM is based on
their similarity to in vivo conditions for cell attachment,
proliferation, and differentiation while maintaining
biomechanical functionality and biocompatibility.
Among biological scaffolds used for clinical application in
regenerative medicine and for current tissue engineering
approaches, decellularized tendon tissue plays a pivotal role.
This is due to a high and still increasing incidence of
tendon pathologies, such as injuries of the Achilles tendon and
traumatic rupture of the anterior cruciate ligament, in an
aging population with growing sporting ambitions [2–4].
Moreover, currently available treatment options for
tendinopathies are often not evidence-based and do not
effectively prevent re-injury, placing a heavy burden on the
health care system and the social economy . Therefore,
tissue engineering involving decellularization techniques
aims not only to prepare 3D-culture models for in vitro
research , but also to produce clinically available
tissue grafts for the reconstruction of musculoskeletal
defects [1, 7] with the ultimate goal to translate tendon
restoration into clinics.
The distinguishing and so far inimitable feature of
site-specific homologous tissue for graft-based tendon
reconstruction is its highly specific ECM composition,
reflecting original biochemical and biomechanical tissue
characteristics at its best. Since synthetic biomaterials
turned out to be more or less inadequate for tendon
reconstruction, biological tendon grafts including auto-,
allo- and xenografts are considered as gold standard for
tendon repair [1, 5]. Especially due to donor site
morbidity, abundantly available allo- and xenografts are favoured
for the development of scaffolds reflecting natural tendon
ECM composition [1, 8]. In order to avoid rejection after
implantation due to cell associated immune response, and
to allow re-seeding procedures, decellularization is an
essential step towards the application of appropriate
tendon-derived scaffolds .
To date, different protocols for decellularization of
tendon tissue of various species have been evaluated,
using physical as well as chemical and/or enzyme-based
methods. However, published protocols for
decellularization of natural tendon tissue provide little insight
regarding the impact of specific decellularization steps
. Physical treatments, like agitation or sonication,
mechanical massage or pressure, or freeze-thaw cycles,
are the most commonly used techniques to disrupt cell
membranes, detach cells within their ECM network and
allow further rinsing to remove cell remnants [5, 10].
Whereas the sole application of freeze-thawing was
reported as insufficient for tendon decellularization ,
Burk et al. reported a significantly more effective
decellularization of large tendon samples by combining repetitive
freeze-thaw cycles with a detergent treatment when
compared to a sole application of detergents . Although
procedural steps such as freeze-thaw cycles are considered
as key procedure, there is no consensus on a certain
protocol or standardized parameters for evaluation of
their effectiveness [12–14].
Considering safety of application in clinical use and the
necessity of commercial graft production under good
manufacturing practice (GMP) conditions, standardization
and accurate documentation of all procedural steps are
crucial. Furthermore, identical processing of large sample
quantities will be required, which can only be achieved by
automating key procedures of decellularization. Therefore,
our study aimed to directly compare the influence of
manual and automated freeze-thaw cycles on the
decellularization effectiveness in equine superficial digital flexor
tendons, based on a previously described decellularization
protocol . Further, we intended to evaluate the impact
of two different cooling as well as heating rates in
automated freeze-thaw cycles with regard to
decellularization effectiveness. To our knowledge, this is the first
description of using automated freeze-thaw cycles for
tissue decellularization, leading to standardization and
optimization of decellularization protocols.
Freshly collected equine superficial digital flexor tendon
samples (n = 10) were used to assess the effectiveness of
automated freeze-thawing as part of an otherwise
established decellularization protocol. Two different
freezethawing protocols were carried out using a controlled
rate freezer (PLANER® Kryo 360 - 1.7). Additionally, a
manual freeze-thawing procedure in accordance with an
established protocol was done.
In all three protocols (Auto-Protocol 1 and 2 and
Manual-Protocol, respectively), after decellularization
was initiated by repetitive freeze-thaw cycles, the same
subsequent procedure of detergent treatment was applied.
Parameters to evaluate the effectiveness of decellularization
included histologically visible nuclei and DNA content. In
addition, further tendon samples (n = 3) were re-seeded
with allogeneic equine adipose tissue-derived mesenchymal
stromal cells (AT-MSC) to evaluate cytocompatibility.
Fresh cadaver limbs from middle-sized warmblood
horses obtained at a local abattoir served as source of
tendon samples. The aseptically performed collection of
tendon specimens from the palmar/plantar aspect of the
mid-metacarpus/-metatarsus was done within 6 h after
slaughter under sterile working conditions. During this
period of time, the skin of the cadaver limbs remained
closed to keep the tendon tissue sterile. To further
prevent bacterial contamination, the recovered tendon
samples were placed in phosphate buffered saline (PBS;
Sigmal Aldrich) supplemented with 2%
penicillinstreptomycin (Sigma Aldrich) and 0.1% gentamycin (Carl
Roth) for overnight storage at 4 °C. Immediately before
automated or manual freeze-thawing, the stored tendon
specimens were washed 5 min each in PBS and 70%
ethanol two times.
From each 8 cm long tendon sample, 2 cm were
separated to serve as internal control, whereas the remaining
6 cm were divided into three equal 2 cm long parts. The
cross-sectional dimension of the tendons were left
unchanged, measuring roughly 1.5 cm × 0.5 cm. Each of
the latter tendon parts underwent decellularization
according to one of three different protocols (Table 1). In
group 1 and 2 (Auto-Protocol 1 and 2), automated
freeze-thaw cycles with controlled cooling and heating
rates differing in the applied temperature change per
minute were performed. Safe and sterile placement of
tendon samples inside the controlled rate freezer was
ensured by using plastic sampling bags (Carl Roth; n = 7
tendon samples) or 15 ml conical centrifuge tubes
(VWR; n = 3 tendon samples). For this purpose, the tube
holders of the controlled rate freezer had been modified
in collaboration with the manufacturer. Group 3 samples
(Manual-Protocol) were subjected to manual
freezethaw cycles including five cycles of 2 min freezing in
liquid nitrogen and 10 min thawing in PBS at 37 °C.
Further decellularization was carried out at room
temperature as well as under continuous agitation and
was performed in the same way for all three groups. For
the purpose of rinsing and induction of cell lysis by
Table 1 Sample groups and decellularization protocols
osmotic effects, all samples were incubated in hypotonic
solution (distilled water) for 48 h. Afterwards, tendon
samples were incubated for 48 h in Tris buffer (Carl
Roth) (pH 7.6) containing 1% Triton X-100 (Carl Roth).
Decellularization was completed by the following
washing steps: 2 × 15 min in distilled water, 24 h in cell
culture medium [DMEM 1 g glucose/L (Thermo Fisher
Scientific) supplemented with 10% fetal bovine serum
(FBS; Sigma Aldrich), 1% penicillin-streptomycin (Sigma
Aldrich), 0.1% gentamycin (Carl Roth)], and again 24 h
in PBS. The performed incubation in cell culture
medium served not only the purpose of rinsing to
remove cellular remnants and residual chemicals, but
also to prepare the scaffolds for cell culture optimally.
Assessment of decellularization effectiveness
Histology and nuclei count
For this analysis, one piece of tendon tissue was obtained
from the centre of each sample. These pieces were fixed
in 4% paraformaldehyde and embedded in paraffin.
Hematoxylin and eosin staining of two longitudinal 6 μm
sections per sample followed. Three randomly chosen
regions of the prepared slides were photographed at 20 ×
magnification (Leica DMi1, Leica MC 170HD, LAS V4.5
Software, Leica Microscope CMS GmbH) and visible cell
nuclei were counted. Results from visible nuclei count
were normalized to the respective internal controls and
are given as percentages relative to controls.
To calculate DNA content, pieces of tendon tissue
recovered from the centre of each sample were subjected
to papain digestion. For this purpose, 200 mg of each
sample were minced into 1 mm3 pieces and washed in
PBS. Subsequently, the samples were incubated in 800 μl
papain digestion buffer [2 mM n-acetyl-l-cysteine (Sigma
Aldrich), 2 mM EDTA (Carl Roth), 50 mM Na2HPO4
(Carl Roth)] and 20 μl papain solution (10 mg/ml)
10 min in 37 °C PBS
2 min in liquid nitrogen
Equine superficial digital flexor tendon samples of group 1 and group 2 were processed by automated freeze-thaw cycles, differing in the performed cooling and heating
rates (Auto-Protocol 1 and Auto-Protocol 2). Both of the applied cooling and heating rates describe a temperature change per unit time. For Auto-Protocol 1 as well as for
Auto-Protocol 2 the maximum reached temperature was + 20 °C (thaw hold for 10 min) and the minimum reached temperature was -80 °C (freeze hold for 3 min). All
temperature regulations of the automated freeze-thaw cycles were carried out by a controlled rate freezer (PLANER® Kryo 360–1.7) that utilizes liquid nitrogen to adjust
temperature. Group 3 included manual freeze-thaw cycles. Further steps of decellularization were the same for all sample groups. Tendon samples classified as internal
control underwent no decellularization
(Sigma Aldrich) for 24 h at 60 °C. Storage of digested
tissue samples before further analysis took place at -20 °C.
DNA content was measured using the Quant-IT™
PicoGreen® dsDNA assay kit (Thermo Fisher Scientific).
To achieve this, digested tissue samples (10-fold dilution
for internal controls) were pipetted in 96-well plates and
an equal volume of PicoGreen® reagent working solution
was added. Subsequently, the samples were incubated
for 5 min at room temperature, protected from light.
Fluorescence (excitation wavelength of 480 nm) was
measured by a microplate reader (SynergyTM H1 Hybrid
Multi-Mode Microplate Reader, Gen5TM Software,
BioTek® Instruments, Inc.). DNA content was
determined by the use of standard curves prepared
from DNA standards measured on the same plates.
Results from DNA measurement were again normalized
to the respective controls and are given as percentages
relative to controls.
Assessment of cytocompatibility
Allogeneic equine AT-MSC were isolated by enzymatic
digestion with collagenase I (Thermo Fisher Scientific)
and expanded until passage 3. Plastic-adherence, trilineage
differentiation potential and expression of MSC-related
surface markers were shown before further processing of
the cells (data not shown). Cells cultured to approximately
60-80% of confluence were detached by trypsinization
and suspended in cell culture medium for re-seeding
Scaffolds with a 2 cm2 surface (2 cm × 1 cm × 2 mm)
from tendons that were decellularized by automated and
manual protocols as described above were manually
prepared using stirrup-shaped blades (Carl Roth). AT-MSC
were then seeded onto the surface of the tendon
scaffolds (130,000 cells in 30 μl / cm2 scaffold surface). After
incubation at 37 °C and 5% CO2 for 4 h, all seeded
tendon scaffolds were covered with cell culture medium
and further incubated for 3 d at 37 °C and 5% CO2.
Histology and LIVE/DEAD® staining
To assess cell morphology and cell integration into the
scaffold, hematoxylin and eosin staining of paraffin
sections of the seeded scaffolds was performed as described
above. Further, LIVE/DEAD® staining of seeded scaffolds
was carried out using the staining kit [LIVE/DEAD®
Viability/Cytotoxicity Kit, for mammalian cells, Thermo
Fisher Scientific; calcein AM 4 mM in anhydrous
DMSO, ethidium homodimer-I 2 mM in DMSO/H2O 1
: 4 (v/v)] according to the manufacturer’s instructions.
The latter was used to evaluate cell viability and
morphology as well as alignment and distribution of the seeded
cells on the scaffold surface.
Statistical data analysis was performed using SPSS®
Statistics 22.0. For comparison of parameters among
different groups, Friedman tests were used. Further analysis
by Wilcoxon signed-rank tests was then carried out for
paired comparisons. As there were no significant
differences between tendon samples that underwent
automated freeze-thaw cycles encased either by the plastic
sampling bags or by the 15 ml centrifuge tubes,
evaluation of decellularization effectiveness was performed
regardless of the sample holder used. The level of
significance was defined at p = 0.05.
Performance of the controlled rate freezer
Graphical records printed by the controller of the
controlled rate freezer plotted the actual measured
temperature profile of Auto-Protocol 1 or 2 versus the
programmed temperature profile (Fig. 1). The controlled
rate freezer repeatably reached the required set point
temperatures within the programmed period of time and
with good accuracy in Auto-Protocol 2. However, in
Auto-Protocol 1, which included faster cooling and
heating rates, a growing delay was repeatedly observed
compared to the programmed temperature profile.
Furthermore, short-term temperature over- or undershoots
were constantly evident at the beginning of each
programmed hold at the required set point temperatures.
This effect was moderate for Auto-Protocol 2 but more
obvious in the records for Auto-Protocol 1.
Effectiveness of decellularization
All of the performed protocols resulted in both reduced
cell nuclei count and reduced DNA content (p = 0.005)
relative to respective controls, with no significant
differences among samples from the three different
freezethaw procedures (Fig. 2).
Histologically, we observed very few visible nuclei
within a uniformly structured ECM (Fig. 3). Empty gaps
were observed, free of cellular residues, between
regularly aligned collagen fibres. In contrast, histologically
evaluated control scaffolds showed a preserved cellular
integrity with rarely occurring signs of apoptosis.
Normalized percentages of residual nuclei in tendon samples
treated with Auto-Protocol 1 (mean value: 2.6%; range:
0–11.9%) and 2 (mean value: 2.5%; range: 0–6.1%) were
slightly higher than those treated manually (mean value:
1.6%; range: 0–4.3%).
Mean values of residual DNA (normalized percentage
values) for all applied protocols fall within a narrow
range of 12.8% (Auto-Protocol 1; range: 9.2–18.2%) and
13.7% (Manual Protocol; range: 8.3–18.9%;
Protocol 2; range: 9.3–16.3%). The obtained data for
Fig. 1 Temperature profiles of Auto-Protocol 1 (group 1) (a) and Auto-Protocol 2 (group 2) (b). Representative graphics for group 1 (a) (Auto-Protocol 1;
cooling/heating rate of 50 °C/min) and for group 2 (b) (Auto-Protocol 2; cooling/heating rate of 20 °C/min). Blue curves represent actual values and brown
curves show target values of the temperature. Both graphics are prepared on the basis of printed temperature records of the biological controlled rate
freezer (PLANER® Kryo 360–1.7) by the use of Adobe® Illustrator® CS6 software
Fig. 2 Visible nuclei count (a) and DNA content (b) of decellularized tendon samples (n = 10). Mean values of residual nuclei count (a) and
residual DNA (b) in % relative to the controls (n = 10). The vertical error bars indicate the confidence interval of 95%. There were no significant
differences in the number of residual nuclei and in the amount of DNA content among tendon samples of both automated protocols (group 1
and 2) and the manually performed protocol (group 3) for decellularization
Fig. 4 Histological assessment of cytocompatibility. Representative images of hematoxylin and eosin stained equine superficial digital flexor tendon
samples after decellularization by automated (a) and manual (b) freeze-thaw cycles, re-seeding with equine adipose tissue-derived mesenchymal
stromal cells and 3 days of culture. A successful re-seeding procedure of the tendon surface is indicated by the dense cell layer adhering to the sample
surface, with a lower number of cells penetrating deeper tissue structures
Fig. 3 Histological assessment of decellularization effectiveness. Representative images of hematoxylin and eosin stained equine superficial digital
flexor tendon samples of group 1 (Auto-Protocol 1) (a), group 2 (Auto-Protocol 2) (b), group 3 (Manual Protocol) (c), showing an apparent
reduction of visible nuclei compared with tendon samples of the internal controls (no decellularization) (d). Decellularized tendon samples of all
groups reveal regularly aligned collagen fibrils and interfibrillar tissue gaps instead of resident cells
decellularized tendon samples correspond to a range
from 9.7 to 12.7 ng DNA/mg wet weight.
Hematoxylin and eosin staining of re-seeded tendon
scaffolds revealed successful seeding on the surface of
automatically and manually processed scaffolds (Fig. 4).
At day 3 after re-seeding, the majority of visible cells
were attached to the scaffold surface, only a very low
number of cells populated deeper tissue layers.
LIVE/DEAD staining of re-seeded scaffolds showed that
the majority of the cells was vital, indicated by green
fluorescence, and a lower number of cells with damaged
membranes indicated by red fluorescence.
Morphologically, vital cells appeared elongated with collagen fibre
oriented alignment, whereas roundly shaped, damaged
cells showed no tendency for directed orientation (Fig. 5).
The observed cell distribution was slightly inhomogeneous.
Besides areas with an even distribution, there were also
only sparsely populated parts of the scaffold surface.
Fig. 5 Fluorescence microscopic assessment of cytocompatibility. Representative panels of LIVE/DEAD® staining of equine superficial digital flexor
tendon specimens decellularized by automated (a) and manual (b) freeze thaw cycles. Decellularized scaffolds were re-seeded with equine adipose
tissue derived mesenchymal stromal cells and a fluorescence microscopic evaluation was performed after 3 days of culture. Vital cells are indicated by
green fluorescence (display of intracellular esterase activity), cells with defect cellular membranes show a red fluorescence signal of their nucleus
The present study demonstrated the same high
effectiveness of both automated and manual freeze-thaw cycles
in decellularization of large tendon samples (equine
superficial digital flexor tendon). Since the sole
application of repeated freeze-thaw cycles for successful
decellularization of natural tendon tissue has been reported
as insufficient [10, 11], all protocols applied in this study
were combined with a Triton X-100-based chemical
treatment. Triton X-100 as a non-ionic detergent is
commonly used in various decellularization protocols to
solubilize cell membranes and dissociate DNA from
proteins . Since a lack of ionic charge results in a
low impact on protein structures, non-ionic detergents
are among the widely used chemical reagents for tissue
decellularization . However, there are conflicting data
about the outcome of the decellularization effectiveness
especially of Triton X-100 in current literature [16–23].
A direct comparison between treatment protocols
particularly for decellularization of tendon tissue is difficult
because of variations in the used concentrations,
combinations and inconsistencies in data analysis and the
times studied. Here, Triton X-100 was mainly chosen
based on the results of our previous study, which had
demonstrated the use of freeze-thaw cycles combined
with this detergent to be effective for decellularization of
full-thickness equine superficial digital flexor tendons
and to maintain scaffold cytocompatibility .
Not only the generally high diversity in applied
protocols for tendon decellularization, but also a lack of
wellstudied decellularization procedures especially for equine
tendon tissue complicate a methodological comparison
with focus on standardization. Referring to
decellularization of equine flexor tendons, there are also successful
decellularization protocols applying ionic detergents
(sodium dodecyl sulfate; SDS) or organic solvents (tributyl
phosphate; TnBP) as chemical agents for decellularization
[23, 24]. But as far as the authors know, beside published
data of Burk et al. there are only very few studies using
repeated freeze-thawing for decellularization of equine
flexor tendons [11, 14]. Moreover, the latter is focused on
viability and biosynthesis of tendon matrix-seeded cells,
rather than on a structured evaluation of decellularization
effectiveness. Finally, to the best of our knowledge, there
is no methodological gold standard for decellularization of
full-thickness equine superficial digital flexor tendons
mentioned in the literature so far.
In the present study, we inferred no significant difference
between automated and manual freeze-thaw procedures
with regard to histologically visible cell nuclei count. Both
procedures resulted in roughly 2% remaining nuclei only.
A direct comparison of histologically assessed residual
nuclei to previous studies is difficult due to different scaffold
size, diverse origins of the decellularized tendon and
ligament tissue samples, and differing procedural steps of
decellularization. The here reported results are roughly in
accordance with a 100% removal of chicken flexor
digitorum profundus cells by using a protocol including Triton
X-100 and peracetic acid (PAA) . Further, Burk et al.
showed a reduction in resident cells of 99% in
decellularization of equine superficial digital flexor tendon samples
when processed by freeze-thawing and Triton X-100 .
However, protocols including Triton X-100 did not result
in a successful removal of cellular remnants and led to a
disrupted tendon structure in rat tail tendons and
central tendon of porcine diaphragms [21, 22]. A recently
published study described decellularization of equine
superficial digital flexor tendon specimens in a
dimension of 10 cm × 1.5 cm × 0.3 cm by using 1% TnBP in
combination with 0, 1, 3 or 5% PAA. The obtained
results showed an increasing reduction in histologically
visible cells (63 to 99%) when treated with 0 to 5%
PAA. However, the use of 5% PAA led to a significant
decrease of proteoglycans and alterations of the ECM,
such as opened collagen fibres as well as an increased
pattern of the tendon crimp .
Further, the present study revealed no significant
differences in DNA-content by direct comparison of
automated and manual protocols, with roughly 13% residual
DNA in all decellularization groups. Thereby, the
presented results are in accordance with current literature
[11, 18, 23]. However, Pridgen et al. showed no
significant reduction in DNA content by using Triton X-100
for decellularization of human flexor tendons . The
latter illustrates again the high diversity in published
data and the resulting difficulties in comparison of
decellularization protocols for tendon tissue.
To our knowledge, this study is the first attempt to partly
automate the process of decellularization of tendon tissue.
Whereas currently published decellularization protocols for
natural tendon tissue describe manually performed
techniques for freeze-thawing steps [12–14, 23, 25–27], our
technique provides a viable alternative to this strategy. Due
to the more standardized and optimized processing with
continuous documentation, automation of repetitive
freezethawing procedures appears as a vital step for the
development of future decellularization protocols.
Usually, controlled rate freezers are utilized in the field
of cryopreservation  and for experimental studies
that evaluate the effects of freezing on diverse tissue and
cell characteristics , focusing on preserving vitality in
terms of storage. Therefore, these machines are designed
for relatively slow cooling procedures, as required for
gentle freezing of cells. This is in accordance with the here
reported temperature profiles of the controlled rate
freezer that showed a more precise course of temperature
for slower cooling rates.
On the contrary, the present study utilized a
conventional liquid nitrogen-based controlled rate freezer for
physical decellularization of natural tendon tissue, for
which rapid freeze-thawing is commonly used. The
desired effect of freezing in terms of decellularization is
a direct cell injury with minimal alteration of ECM
composition. Since all steps of freeze-thawing, including
cooling rate as well as the coldest set point temperature,
freezing hold time, thawing rate and number of
repetitions, are able to influence cell injury , the present
study applied two different cooling and heating rates.
Their direct comparison revealed no significant
difference in their effectiveness for decellularization of tendon
samples. A slow freezing rate, associated with
extracellular ice crystal formation and a subsequent hyperosmolar
shift in the extracellular environment, leads to cell
dehydration as well as intracellular solute concentration,
which is not always lethal for resident cells. Conversely,
faster cooling rates are associated with intracellular ice
crystal formation, leading almost always to cell death
due to disintegrated organelles and cell membranes .
However, the obtained data suggest a minor importance
of the applied cooling rate for decellularization
effectiveness in large tendon samples. For further interpretation
and especially for a comparison to
freeze-thawingassociated effects in other connective tissue types, it
should be noted that response to and threshold for
cooling rates are cell-specific features [31, 32]. Moreover, the
disrupting effects of intracellular ice crystals depend on
a sufficient duration of thawing . Finally, in this
study, the controlled rate freezer accurately performed
slow cooling and heating rates of -20 °C per min and
+20 °C per min. As this procedure was effective for
decellularization of large tendon samples, the utilized
controlled rate freezer is suitable for this purpose. A
possible methodological refinement is related to the
requirement of liquid nitrogen. Given the problematic risk
of contamination during transport or storage, the use of
liquid nitrogen under GMP conditions is limited .
Therefore, future investigations aiming at a more
extensive sample processing may benefit from alternative
strategies providing cryogenic temperatures.
For further characterization of the decellularized tendon
scaffolds, the present study provided a basic assessment of
cytocompatibility by re-seeding the tendon surface with
equine AT-MSC. In the field of tissue engineering,
reseeding procedures of decellularized tendon matrices are
still challenging since their dense structure makes a
satisfactory cell infiltration into deeper tissue layers
complicated . Nevertheless, promising results of cell seeded
scaffolds for tendon regeneration justify the search for
more appropriate re-seeding techniques [33–36]. In the
present study, tendon samples that underwent automated
as well as manual freeze-thaw cycles could successfully be
re-seeded. However, as reported before, only a very low
number of cells penetrated deeper tissue layers,
emphasizing the need for improved re-seeding techniques. The
latter should aim for residual-free tendon scaffolds with a
loosened matrix to allow ingrowth of scaffold-seeded cells
into an almost completely preserved tissue-specific ECM.
Among already published technical approaches
promising effects resulted especially from using ultrasound
sonication, PAA treatment and several injection
techniques [1, 8, 37–39]. Further, the reduction in scaffold
dimension to a thickness of 300 μm led to an improved
tissue penetration of cells in the present experimental
set-up (unpublished data) as well as in already published
studies . Although the majority of cells seeded on the
tendon scaffolds was vital, there were also roundly shaped,
damaged cells observed by LIVE/DEAD staining in the
present study. Referring to possible causes that led to
damaged and detached cells, future re-seeding procedures
should reduce manual handling of the seeded constructs.
Other factors that mainly influence cell viability include
chemical residues with cytotoxic effects and the initial cell
seeding density. Particularly, metabolic stressors such as
nutrient availability and metabolic byproducts of cellular
origin gain in importance as cell density increases .
The horse is referred to as the fourth most frequently
used source of tendon tissue for decellularization . In
terms of translation, the equine superficial digital flexor
tendon is considered to be particularly important since
its functional, structural, and pathological characteristics
have already been subject of extensive research [41, 42]
and are similar to those of the human Achilles tendon.
Therefore, the horse is considered as the ideal animal
model for human tendon pathologies, such as
exerciseinduced Achilles tendon injury . With regard to
providing commercially available tissue grafts for
xenotransplantation, equine tendon tissue offers further
benefits. Those include the accessibility related to
breeding and slaughter, the provided dimension of tendon
tissue required in reconstructive surgery, and the limited
number of zoonotic agents .
Future studies should aim to assess further characteristics
of tendon matrices that underwent automated freeze-thaw
cycles. These could include an evaluation of ECM
ultrastructure and biochemical composition, of mechanical
properties, of antigen removal and of adverse host reactions
induced by the tendon scaffolds. To allow the production
of commercially available tendon graft materials for use in
human as well as in veterinary medicine, high quality
manufacturing processes are required. Therefore,
automation of so far manually performed key procedures in tissue
decellularization, such as freeze-thawing, is essential.
Moreover, future methodological research should focus on the
automation of further decellularization steps. The latter
especially include automated washing steps under
continuous agitation performed in a closed system and applicable
for a high number of samples. Finally, increased
requirements with regard to validation, reproducibility, and safety,
need to match with practicable approaches.
Automated freeze-thaw cycles carried out by a liquid
nitrogen-based controlled rate freezer are effective for
decellularization of large tendons when combined with a
non-ionic detergent treatment (Triton X-100). These
findings are an essential step towards a standardized
production of decellularized tendon scaffolds for various in
vitro applications and further development of graft-based
reconstruction of musculoskeletal defects.
AT - MSC: Adipose tissue-derived mesenchymal stromal cells;
ECM: Extracellular matrix; FBS: Fetal bovine serum; GMP: Good manufacturing
practice; PAA: Peracetic acid; PBS: Phosphate buffered saline; SDS: Sodium
dodecyl sulfate; TnBP: Tributyl phosphate
The work presented in this paper was made possible by funding from the
German Federal Ministry of Education and Research (BMBF 1315883), the
German National Research Foundation and the Saxon Ministry of Science
and the Fine Arts. We acknowledge support from the German Research
Foundation (DFG) and Universität Leipzig within the program of Open
SMG carried out the experiments, analysed the data and helped to draft the
manuscript. AA, IE, SH helped to design the study. JB designed the study,
performed the statistical analysis of the data and revised the manuscript. SPR
helped to perform manual and automated decellularization, analysed the data,
and wrote the manuscript. All authors read and approved the final version of
Ethics approval and consent to participate
All of the donor animals were sacrificed for reasons unrelated to the current
study. Therefore, in accordance with national guidelines and the local ethics
committee (Landesdirektion Leipzig), no license was required for sample
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