![A representation of inkjet bioprinting of droplets onto collection plates with a piezoelectric actuator (a) and a thermal actuator (b) [38].](https://www.researchgate.net/publication/356171725/figure/fig1/AS:1089333150916608@1636728670704/A-representation-of-inkjet-bioprinting-of-droplets-onto-collection-plates-with-a_Q320.jpg)
![A representation of laser-assisted bioprinting with bioink droplets deposited by laser pulses onto a collector slide [38].](https://www.researchgate.net/publication/356171725/figure/fig2/AS:1089333150920709@1636728670740/A-representation-of-laser-assisted-bioprinting-with-bioink-droplets-deposited-by-laser_Q320.jpg)
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sensors
Review
Bioprinting of Stem Cells in Multimaterial Scaffolds and Their
Applications in Bone Tissue Engineering
Shebin Tharakan 1,2, Shams Khondkar 1,3 and Azhar Ilyas 1, 4, *
Citation: Tharakan, S.; Khondkar, S.;
Ilyas, A. Bioprinting of Stem Cells in
Multimaterial Scaffolds and Their
Applications in Bone Tissue
Engineering. Sensors 2021,21, 7477.
https://doi.org/10.3390/s21227477
Academic Editor: Rawil Fakhrullin
Received: 24 August 2021
Accepted: 5 November 2021
Published: 10 November 2021
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Copyright: © 2021 by the authors.
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conditions of the Creative Commons
Attribution (CC BY) license (https://
creativecommons.org/licenses/by/
4.0/).
1Bio-Nanotechnology and Biomaterials (BNB) Lab, New York Institute of Technology, Old Westbury,
NY 11568, USA; stharaka@nyit.edu (S.T.); skhondka@nyit.edu (S.K.)
2New York Institute of Technology, College of Osteopathic Medicine, Old Westbury, NY 11568, USA
3Department of Bioengineering, New York Institute of Technology, Old Westbury, NY 11568, USA
4Department of Electrical and Computer Engineering, New York Institute of Technology, Old Westbury,
NY 11568, USA
*Correspondence: ailyas@nyit.edu
Abstract:
Bioprinting stem cells into three-dimensional (3D) scaffolds has emerged as a new avenue
for regenerative medicine, bone tissue engineering, and biosensor manufacturing in recent years.
Mesenchymal stem cells, such as adipose-derived and bone-marrow-derived stem cells, are capable
of multipotent differentiation in a 3D culture. The use of different printing methods results in
varying effects on the bioprinted stem cells with the appearance of no general adverse effects.
Specifically, extrusion, inkjet, and laser-assisted bioprinting are three methods that impact stem cell
viability, proliferation, and differentiation potential. Each printing method confers advantages and
disadvantages that directly influence cellular behavior. Additionally, the acquisition of 3D bioprinters
has become more prominent with innovative technology and affordability. With accessible technology,
custom 3D bioprinters with capabilities to print high-performance bioinks are used for biosensor
fabrication. Such 3D printed biosensors are used to control conductivity and electrical transmission
in physiological environments. Once printed, the scaffolds containing the aforementioned stem
cells have a significant impact on cellular behavior and differentiation. Natural polymer hydrogels
and natural composites can impact osteogenic differentiation with some inducing chondrogenesis.
Further studies have shown enhanced osteogenesis using cell-laden scaffolds
in vivo
. Furthermore,
selective use of biomaterials can directly influence cell fate and the quantity of osteogenesis. This
review evaluates the impact of extrusion, inkjet, and laser-assisted bioprinting on adipose-derived
and bone-marrow-derived stem cells along with the effect of incorporating these stem cells into
natural and composite biomaterials.
Keywords: bioprinting; stem cells; composite biomaterials; osteogenesis; fracture repair
1. Introduction
Bone fractures in the United States are projected to increase 50% by 2025. Individuals
in the age group 65 to 74 are estimated to have the fastest increase of 87% [
1
]. The
increasing rate of fractures warrants novel and innovative methods of treatment. Autografts
and allografts are the standard clinical solutions, with their respective advantages and
disadvantages. Bone autografts are continuously considered the orthopedic gold standard
in bone tissue transplantations. Autologous bone grafts are commonly taken from the iliac
crest with donor site morbidity associated with the transplant [
2
]. New studies, however,
are examining the potential efficacy of proximal tibial grafts due to lower post-operative
pain and similar healing properties to the iliac crest [
3
,
4
]. Autografts are resorbable,
osteoconductive, osteoinductive, and provide a living source of cells [
5
]. Donor site
morbidity and infection are common concerns regarding autografts [
6
]. Unlike autografts,
allografts require the graft from a cadaver or another individual. Cadavers lack living
osteoblasts and osteoprogenitors, thereby limiting the osteogenic potential of the graft [
7
,
8
].
Sensors 2021,21, 7477. https://doi.org/10.3390/s21227477 https://www.mdpi.com/journal/sensors
Sensors 2021,21, 7477 2 of 20
Additionally, allografts are associated with a greater risk of an immune response and graft
rejection but have lower donor site morbidity [
5
]. Apart from autologous or allograft bone
transplantations, synthetic or natural polymeric materials may be used in their place.
Synthetic or natural polymers may be used in the place of autografts. These polymers
need to be comparable to autografts, and with fewer disadvantages, in order to be used
effectively in a clinical setting [
9
]. The assembly of synthetic or natural materials into
usable and implantable constructs can be achieved with 3D bioprinting to create scaffolds.
3D bioprinting is an extension of additive manufacturing. 3D bioprinting, by definition, is
the printing of constructs with the incorporation of viable cells, biomaterials, or biological
materials [
10
,
11
]. The initial chemical mixture is referred to as the bioink, and once bio-
printed it becomes known as a scaffold or construct, and can be implanted. With recent
advances in 3D bioprinting technology, biomimetic scaffolds are used to accelerate bone
regeneration
in vitro
and
in vivo
using hydrogels or composite structures. Hydrogels are
a gel-like macromolecular complex creating a 3D network of polymers, with polymers
such as collagen, hyaluronic acid, or alginate. Bone is traditionally considered dense
solid tissue, with the principal component being hydroxyapatite. Thus, repair of bone
can be associated with metal implants such as titanium [
12
]. In contrast, hydrogels are
soft gel-like constructs that can be directly applied to bone to induce regeneration with
the inclusion of cells and growth factors. The efficacy of soft materials has been shown
through
in vivo
calvaria and femoral defect models, while clinically a mix of both soft and
hard materials such as metals can be used. Additionally, hydrogels can be 3D bioprinted
as composites, meaning that they can contain multiple polymers intertwined to influence
the physiochemical properties of the printed scaffold. The hydrogel scaffolds implement
natural or synthetic biomaterials to form porous constructs to repair fractures in place
of an autograft [
13
]. An ideal scaffold is biocompatible, biodegradable, osteoconductive,
osteoinductive, and can resist compressive forces. Natural materials include proteins such
as collagen, silk, and fibrin and polysaccharides such as alginate, chitosan, and hyaluronic
acid. Natural polymers allow cell incorporation with high cell viability but have disad-
vantages such as low mechanical strength and varying biodegradability. The use of these
materials creates natural scaffolds once the bioink has undergone 3D bioprinting. Natural
polymers tend to have high cell adhesion due to the presence of cell adhesion molecules
such as integrins [
14
,
15
]. A notable exception is alginate which lacks these binding sites
for cells. This is resolved by functionalizing alginate with a RGD peptide sequence to
permit cell adhesion [
16
]. Synthetic polymers are the opposite with increased mechanical
strength and greater fine-tuning of biodegradability, forming synthetic scaffolds [
17
]. Un-
like natural polymers, synthetic materials are lacking in bioactivity [
18
]. Included in this
class of polymers are poly(
ε
-caprolactone) (PCL), poly(lactide-co-glycolide) (PLGA), and
polyglycolic acid (PGA) [
19
]. Alternatively, functionalized polymers can be used in place
to demonstrate similar efficacy. Multiple materials can be combined to form a composite
scaffold which assimilates the properties of each polymer. These combinations confer
improved biocompatibility, biodegradation, and mechanical properties to improve the
desired parameters [20–22].
The hydrogel constructs can be printed with cells such as osteoblasts or metal ions to
further speed up the healing process [
21
,
22
]. Alternatively, mesenchymal stem cells can be
used in place of osteoprogenitor or osteoblast cells. Mesenchymal stem cells (MSCs) are
multipotent and can differentiate into cartilage, bone, adipose, muscle and other tissues
depending on the growth factors present making them versatile and useful for tissue
engineering. Human mesenchymal stem cells (hMSCs) were initially harvested from bone
marrow, but have now been isolated from adipose tissue, amniotic fluid, placental tissue,
Wharton’s jelly, endometrium, and dental pulp [
23
,
24
]. Li et al. evaluated the osteogenicity
of hMSCs and found Wharton’s Jelly MSCs to have the greatest osteogenic potential, fol-
lowed by placental, adipose, and bone marrow stem cells [
25
]. Adipose and bone marrow
stem cells both have similar osteogenic capabilities, but different disadvantages with their
use. Adipose-derived stem cells (ADSC) are easy to harvest, but require more testing to
Sensors 2021,21, 7477 3 of 20
evaluate their capabilities in bone regeneration, while bone marrow stem cells (BMSC) are
extracted in low quantities and require extensive culturing [
26
]. Coupled with 3D bioprint-
ing, these cells provide osteoinductive capabilities which improve bone regeneration [27].
A benefit of 3D bioprinting with stem cells or cell-lines is the incorporation of cells directly
into the bioink for immediate printing. Compared to seeding cells post printing, 3D bio-
printing cells in conjunction with the biomaterials offers a streamlined process to generate
multiple samples without the waiting time for cell attachment by seeding. A significant
advantage, however, is the homogenous distribution of cells during the printing process,
which may not be conferred during cell seeding. Homogenous dispersion provides the
benefit of a functional culture that can increase the formation of tissue [28]. However, cell
viability must be confirmed post printing due to pressure differentials and stress during
the printing process. Aside from cell stress, some 3D bioprinters are expensive and may
not be academically available.
3D bioprinters are multifaceted and can have uses in different fields, ranging from
tissue engineering to biosensor manufacturing. In particular, 3D bioprinters are capable
of printing high-performance bioink for biosensor applications [
29
]. High-performance
bioinks are next-generation bioinks with reinforcement mechanisms to drive cell func-
tions [
30
]. The functionality of biosensors involves appropriate conductivity and electrical
transmission. Organ-wise, this applies to cardiac tissue due to electrical conduction through
intercalated discs. In contrast to bone tissue, electrical conductivity is not a primary concern
for fracture studies. A recent study on cardiac tissue regeneration involved the use of gold
nanocomposite bioinks to improve cardiomyocyte and cardiac fibroblast functionality [
31
].
The functional use of this bioink can be further developed to create biosensors and medical
devices to assist cardiac function. Furthermore, to demonstrate the diverse potential of 3D
bioprinting, a separate study devised a high-performance alginate bioink with conductive
silver nanoparticles and chondrocytes to synthesize a bionic ear [32]. The potential to use
3D bioprinting to create electrical configurations and biosensors is untold and can only
expand in the future. This paper will expand on three 3D bioprinting techniques and their
impact on the viability, differentiation, and osteogenicity of adipose-derived and bone
marrow stem cells. The osteogenic effects of both stem cells will be evaluated in hydrogel
and composite biomaterial scaffolds to determine their efficacy in bone fracture repair.
2. Bioprinting Techniques
2.1. Extrusion Bioprinting
Extrusion bioprinting is cost effective and is the most used method in 3D bioprinting.
Extrusion bioprinters can contain multiple printheads to mix many materials together
providing various 3D structures comprising of different cells, biomaterials, and signaling
molecules [
33
]. In this method, cartridges are loaded with the biomaterials or bioink
and continuous filaments are printed directly onto a stage (Figure 1). Continuous bioink
filaments instead of droplets are extruded through the printer due to the continuous
force applied. The extrusion mechanism can be pneumatic, piston, or screw driven, with
temperature or pressure being computer controlled [
34
]. Previous studies have indicated
the range of viscosity for printable biomaterials is between 30–6
×
10
7
mPa
·
s indicating
the extrusion method can print very viscous bioinks. The printing resolution is poor, with
the range being between 200–1000
µ
m, compared to other methods. Extrusion fabrication
speed is the slowest, and is between 10
µ
m s
−1
–700 mm s
−1
[
35
]. Current extrusion
bioprinters are capable of printing high cell concentrations, including cell spheroids, but
are limited by lower cell viability due to the high extrusion and shear pressure [36,37].
Sensors 2021,21, 7477 4 of 20
Sensors 2021, 21, x FOR PEER REVIEW 4 of 20
Figure 1. A representation of extrusion bioprinting of bioink onto a collection plate as a continuous
filament [38].
2.1.1. Extrusion Bioprinting of Adipose-Derived Stem Cells
Shear forces have been shown to impact differentiation towards endothelial and bone
lineage [39]. Zhang et al. exposed human ADSCs (hADSC) to 12 dynes of shear stress,
which stimulated endothelial differentiation and acquisition of a nonthrombogenic phe-
notype [40]. Colle et al. printed spheroid ADSCs onto GelMA scaffolds and demonstrated
cell viability was equal at strut edges and centers. After 7 days of culturing the GelMA-
encapsulated spheroids, the viability of the 3D printed scaffolds was 79%, while non-
printed had 80% viability. The printed scaffolds and non-printed scaffolds had no signif-
icant difference in viability [41]. Interestingly, Leucht et al. created and printed a bone
vascularization bioink with hADSCs and human dermal microvascular endothelial cells
(HDMECs). The co-cultured hydrogels consisted of the osteogenic hADSCs and vascular
compartment (hADSCs + HDMECs). The cells were viable after being printed and in-
creased levels of Col I, fibronectin, ALP, and OPN at 20 days in the co-culture demon-
strated osteogenic differentiation [42]. Wang et al. showed that cell viability on day 1 after
printing was 88.13%, with viability increasing to 90.41% 7 days later. The increased ex-
pression of vinculin 7 days after printing showed that cells could attach to the hydrogels
and grow normally. Blood vessel ingrowth and bone matrix formation were observed af-
ter 8 weeks of implantation in vivo, showing osteogenic capacity of hADSC in forming
new bone [43]. The hADSCs pre-differentiated for 3 weeks and printed in a Fibrin/Gela-
tin/Hyaluronic Acid/Glycerol hydrogel demonstrated greater hydrogel calcification than
non-differentiated hADSCs. 3D-printed hADSC monolayers displayed high viability 2
days after printing, with 97.1% of the cells surviving [44]. Extrusion bioprinting ADSCs
provides adequate viability, proliferation, and retention of differentiation ability. The ef-
fect of extrusion printing on adipose-derived stem cells is an avenue for investigation for
bone repair as it has demonstrated potential capabilities in osteogenesis.
2.1.2. Extrusion Bioprinting of Bone-Marrow-Derived Stem Cells
The high shear stress from extrusion bioprinting can induce cells into a certain line-
age. Yourek et al. determined the effects of fluid shear stress on human BMSCs (hBMSC).
The findings suggested that shear stress encourages differentiation into the osteoblast
Figure 1. A representation of extrusion bioprinting of bioink onto a collection plate as a continuous
filament [38].
2.1.1. Extrusion Bioprinting of Adipose-Derived Stem Cells
Shear forces have been shown to impact differentiation towards endothelial and
bone lineage [
39
]. Zhang et al. exposed human ADSCs (hADSC) to 12 dynes of shear
stress, which stimulated endothelial differentiation and acquisition of a nonthrombogenic
phenotype [
40
]. Colle et al. printed spheroid ADSCs onto GelMA scaffolds and demon-
strated cell viability was equal at strut edges and centers. After 7 days of culturing the
GelMA-encapsulated spheroids, the viability of the 3D printed scaffolds was 79%, while
non-printed had 80% viability. The printed scaffolds and non-printed scaffolds had no
significant difference in viability [
41
]. Interestingly, Leucht et al. created and printed a
bone vascularization bioink with hADSCs and human dermal microvascular endothelial
cells (HDMECs). The co-cultured hydrogels consisted of the osteogenic hADSCs and
vascular compartment (hADSCs + HDMECs). The cells were viable after being printed
and increased levels of Col I, fibronectin, ALP, and OPN at 20 days in the co-culture
demonstrated osteogenic differentiation [
42
]. Wang et al. showed that cell viability on
day 1 after printing was 88.13%, with viability increasing to 90.41% 7 days later. The
increased expression of vinculin 7 days after printing showed that cells could attach to the
hydrogels and grow normally. Blood vessel ingrowth and bone matrix formation were
observed after 8 weeks of implantation
in vivo
, showing osteogenic capacity of hADSC
in forming new bone [
43
]. The hADSCs pre-differentiated for 3 weeks and printed in a
Fibrin/Gelatin/Hyaluronic Acid/Glycerol hydrogel demonstrated greater hydrogel calcifi-
cation than non-differentiated hADSCs. 3D-printed hADSC monolayers displayed high
viability 2 days after printing, with 97.1% of the cells surviving [
44
]. Extrusion bioprinting
ADSCs provides adequate viability, proliferation, and retention of differentiation ability.
The effect of extrusion printing on adipose-derived stem cells is an avenue for investigation
for bone repair as it has demonstrated potential capabilities in osteogenesis.
2.1.2. Extrusion Bioprinting of Bone-Marrow-Derived Stem Cells
The high shear stress from extrusion bioprinting can induce cells into a certain lineage.
Yourek et al. determined the effects of fluid shear stress on human BMSCs (hBMSC). The
findings suggested that shear stress encourages differentiation into the osteoblast lineage.
The upregulation of BMP-2, Bone sialoprotein, and Osteopontin after 4 days indicates that
shear stress encourages osteogenic gene expression [
45
]. However, goat BMSCs (gBMSC)
bioprinted by Fedorovich et al. expressed ALP after 2 weeks of culturing in osteogenic
media, implying that stem cells retain their long-term differentiation potential. It was de-
termined that needle diameter had no significant effect on cell viability 5 h after deposition.
Sensors 2021,21, 7477 5 of 20
Bioprinting had no adverse effect on the gBMSCs, but hydrogel composition impacted
cell viability. Matrigel and alginate scaffolds were shown to have greater cell survival
after 7 days compared to agarose and Lutrol F127 scaffolds [
46
]. Rat BMSC (rBMSC)
microbeads printed by pneumatic extrusion appeared morphologically round and were
evenly distributed throughout the alginate dialdehyde-gelatin (ADA-GEL) and nano-scale
glass bead (ADA-GEL-nBG) scaffold. Pneumatic pressure changes from 2.3 to 2.5 bars
during microbead printing had no impact on rBMSC survivability. Cell viability was
85% for the ADA-GEL scaffold and 75% for the ADA-GEL-nBG after 7 days of culturing,
implying that the materials are not cytotoxic [
47
]. The lower viability may be due to the
pneumatic extrusion printing as this method places pressure on the cells. Du et al. printed
methacrylamide gelatin scaffolds with rBMSCs and determined cell viability and DNA
content after mechanical extrusion. Cell viability was 91.8%, but increased to 94.9% on day
28. After extrusion, DNA content was less than 30%, showing immediate low proliferation.
However, by day 28, DNA content increased to almost 70%, indicating proper prolifera-
tion [
48
]. Finally, extrusion bioprinting may have effects on cell survivability and stemness
due to the high shear pressure but generally, no adverse effects appear to occur.
2.2. Inkjet Bioprinting
Unlike extrusion bioprinting, inkjet bioprinting employs discrete droplets as the
primary structural component in 3D constructs deposited onto a collection plate. A thermal
or piezoelectric actuator is used to generate droplets of the desired size by creating pressure
increases to cause propulsion of the bioink (Figure 2). Inkjet bioprinting can be classified
as continuous inkjet (CIJ) bioprinting or drop-on-demand (DOD) bioprinting, with DOD
being the most suitable option for tissue engineering [
49
]. The use of inkjet bioprinting
confers high printing precision, along with low cost and accessibility [
38
,
49
]. However, the
incorporation of viscous cell-laden bioinks can damage the nozzle due to clogging, which
in turn hinders cell viability and function. It has been reported that the printable viscosity
is less than 10 mPa
·
s. Additionally, this printing method has a high resolution of between
10 and 50
µ
m, and a fast fabrication speed of 10
5
droplets/s [
29
,
35
]. The downside of this
method is the inability to print high cell concentrations, since higher cell densities increase
bioink viscosity [
50
]. The reported cell densities are less than 10
6
cells/mL, limiting the
potential for bioprinting with highly viscous biomaterials [35].
Sensors 2021, 21, x FOR PEER REVIEW 6 of 20
piezoelectric actuators on ADSC survivability should be evaluated in-depth. Currently,
ADSCs can differentiate into osteoblasts with the help of osteogenic differentiation media
under hydrostatic pressure [52]. It has also been shown that ADSCs are able to lean toward
a chondrogenic phenotype with no exposure to chondrogenic soluble factors under hy-
drostatic pressure [53]. The effect of inkjet pressure should be explored further to evaluate
the differentiation ability of ADSCs into the osteoblast lineage.
2.2.2. Inkjet Bioprinting of Bone-Marrow-Derived Stem Cells
Blaeser et al. extracted hMSCs from the femoral head of five donors and evaluated
shear stress in cell-laden alginate scaffolds. Cell proliferation and viability were not sig-
nificantly affected at low shear pressures (<5 kPa), but were strongly affected at higher
shear pressures (>10 kPa). Interestingly, medium shear pressures (5–10 kPa) encouraged
cell proliferation, indicating that moderate shear pressure has stimulatory effects [37]. Pre-
viously, high mechanical pressure has been shown to differentiate mesenchymal stem
cells towards an osteoblast lineage [54]. However, the stem cell phenotype remained un-
changed post printing, with the detection of vimentin, a surface marker in mesenchymal
stem cells. The pressure threshold is near 5 kPa for cells to be printed without side-effects
[37]. Gao et al. inkjet bioprinted acrylated peptides and PEG hydrogels with hBMSCs and
determined that cell viability after 24 h was 87.9%, indicating that cells were preserved
post printing. Osteogenic differentiation did not seem to be affected by printing; however,
RUNX2 expression was consistently elevated in the PEG-peptide scaffold, indicating long-
term osteogenic differentiation. ALP levels were markedly increased by day 7, showing
accelerated osteoblast formation [55]. hBMSCs printed in PEG-GelMA scaffolds exhibited
viability greater than 80% immediately after printing. The inkjet printing permitted the
formation of evenly distributed cells in a layer-by-layer fashion for bone tissue synthesis.
Unfortunately, GelMA is highly viscous, which hinders printability. Moreover, the scaf-
folds were simultaneously photopolymerized, which had no significant negative effects
on the hBMSCs [56].
Since this method of printing results in low resolution and cell concentrations,
BMSCs should be printed in low concentrations to avoid nozzle clogging and adverse cell
effects. Optimal bioinks should be developed to maximize BMSC viability, proliferation,
and differentiation to increase their osteogenic effects. Additionally, more studies should
be conducted to evaluate the effects of the inkjet actuators (thermal vs. piezoelectric) on
BMSC stemness, osteogenesis, and viability.
Figure 2.
A representation of inkjet bioprinting of droplets onto collection plates with a piezoelectric actuator (
a
) and a
thermal actuator (b) [38].
Sensors 2021,21, 7477 6 of 20
2.2.1. Inkjet Bioprinting of Adipose-Derived Stem Cells
Kim et al. used piezoelectric inkjet printing to construct an hADSC-laden PLGA
scaffold on polystyrene substrate. The PLGA inks were prepared by dissolving PLGA in
N,N-dimethylformamide followed by 0.2-micron filtration for piezoelectric inkjet printing.
The PLGA printed patterns were favorable for hADSC adhesion, with adhesion >20%, but
the polystyrene was less than 10% 24 h after printing. The choice of biomaterial impacts
immediate cell adhesion, as demonstrated with the greater PLGA cell adhesion. At 24 h
after printing, cell proliferation was 154%. Proliferation rate increased 275% after 72 h post
printing, indicating that the hADSCs retained their proliferative capabilities. Since inkjet
printers are limited to printing lower cell densities, the PLGA patterns were incomplete,
with the hADSCs resulting in partially filled constructs. The variable patterns created by
inkjet printing are simple and suitable for analyzing geometrical effects on hADSC or stem
cell behavior [51].
More studies should be conducted on ADSC phenotype expression, differentiation
potential, and viability immediately after inkjet printing. The effect of the thermal or
piezoelectric actuators on ADSC survivability should be evaluated in-depth. Currently,
ADSCs can differentiate into osteoblasts with the help of osteogenic differentiation media
under hydrostatic pressure [
52
]. It has also been shown that ADSCs are able to lean
toward a chondrogenic phenotype with no exposure to chondrogenic soluble factors under
hydrostatic pressure [
53
]. The effect of inkjet pressure should be explored further to
evaluate the differentiation ability of ADSCs into the osteoblast lineage.
2.2.2. Inkjet Bioprinting of Bone-Marrow-Derived Stem Cells
Blaeser et al. extracted hMSCs from the femoral head of five donors and evaluated
shear stress in cell-laden alginate scaffolds. Cell proliferation and viability were not
significantly affected at low shear pressures (<5 kPa), but were strongly affected at higher
shear pressures (>10 kPa). Interestingly, medium shear pressures (5–10 kPa) encouraged
cell proliferation, indicating that moderate shear pressure has stimulatory effects [
37
].
Previously, high mechanical pressure has been shown to differentiate mesenchymal stem
cells towards an osteoblast lineage [
54
]. However, the stem cell phenotype remained
unchanged post printing, with the detection of vimentin, a surface marker in mesenchymal
stem cells. The pressure threshold is near 5 kPa for cells to be printed without side-
effects [
37
]. Gao et al. inkjet bioprinted acrylated peptides and PEG hydrogels with hBMSCs
and determined that cell viability after 24 h was 87.9%, indicating that cells were preserved
post printing. Osteogenic differentiation did not seem to be affected by printing; however,
RUNX2 expression was consistently elevated in the PEG-peptide scaffold, indicating long-
term osteogenic differentiation. ALP levels were markedly increased by day 7, showing
accelerated osteoblast formation [
55
]. hBMSCs printed in PEG-GelMA scaffolds exhibited
viability greater than 80% immediately after printing. The inkjet printing permitted the
formation of evenly distributed cells in a layer-by-layer fashion for bone tissue synthesis.
Unfortunately, GelMA is highly viscous, which hinders printability. Moreover, the scaffolds
were simultaneously photopolymerized, which had no significant negative effects on the
hBMSCs [56].
Since this method of printing results in low resolution and cell concentrations, BMSCs
should be printed in low concentrations to avoid nozzle clogging and adverse cell effects.
Optimal bioinks should be developed to maximize BMSC viability, proliferation, and
differentiation to increase their osteogenic effects. Additionally, more studies should be
conducted to evaluate the effects of the inkjet actuators (thermal vs. piezoelectric) on BMSC
stemness, osteogenesis, and viability.
2.3. Laser-Assisted Bioprinting
Laser-assisted bioprinting, or laser-induced forward transfer (LIFT), uses droplet
release like the inkjet bioprinting system. A laser pulse encounters the top donor layer,
which forms a bubble to propel the bottom bioink layer as droplets onto the collection plate
Sensors 2021,21, 7477 7 of 20
(Figure 3) [
57
]. Unlike extrusion and inkjet bioprinting, there is no contact with a nozzle,
which eliminates the possibility of clogging and shear pressure. Due to this, cell viability is
higher compared to the other two methods (>95%) and viscous bioinks are printable [
58
].
Printable bioink cell densities are less than 10
8
cells/mL, with viscosity between 1 and
300 mPa
·
s [
35
]. The greatest strength of laser-assisted bioprinting is the high printing speed
and precision, which allows for fine-tuned 3D structures capable of mimicking natural
tissues [
34
]. Printing resolution is reported to be between 10 and 100
µ
m, with fabrication
speed being 200–1600 mm s
−1
[
35
]. This method is the most expensive and complex, which
limits its use commercially [
38
]. However, LIFT is used much more often in bioprinting
and offers potential in printing stem cells due to the capability of creating complex 3D
structures. A comparison of each printing method is summarized in Table 1, which is
adapted from [35].
Sensors 2021, 21, x FOR PEER REVIEW 8 of 20
al. determined that there was no significant difference in apoptosis, proliferation, and gen-
otoxicity in hBMSCs post printing with a Nd:YAG-laser. The hBMSCs demonstrated a
survival rate of 90% after printing [59]. Laser bioprinting confers printing with high reso-
lution and precision, as shown by a printing resolution of 138 µm and precision of 16 µm
in one study with BMSCs [66]. Ali et al. used slow jet conditions, which are more stable,
to minimize droplet impact energy with mice BMSCs (mBMSC). The slow jetting condi-
tions have decreased laser pulse energy, which reduces shear stress. The mBMSCs were
printed with high cell viability, which was measured 24 h after printing, and possessed
high resolution [67]. Laser bioprinting can be used to deposit BMSCs directly in vivo to
enhance osteogenesis. Keriquel et al. devised a method to print nano-Hydroxyapatite
(nHA) layers directly onto a mouse critical sized calvaria defect. The experimentation
demonstrated laser exposure to the dura mater caused temporary inflammation and no
permanent tissue damage in mouse brain [68]. This was further expanded by printing
BMSCs in situ in a ring or disk geometry to induce osteogenesis in vivo. The in situ printed
BMSC nHA disks showed significant osteogenesis than the ring shaped BMSC nHA. It is
hypothesized that due to the disk cell homogeny and proximity, the BMSCs secreted para-
crine factors to induce osteogenic differentiation [69]. This novel technique should be ex-
plored in greater depth with different biomaterials and BMSCs to gauge its full potential.
A summary of each bioprinting technique and its effects on ADSCs and BMSCs is pro-
vided in Table 2.
Figure 3. A representation of laser-assisted bioprinting with bioink droplets deposited by laser pulses onto a collector slide
[38].
Table 1. Bioprinting techniques.
Extrusion [50,70–80] Inkjet [50,51,80–86] Laser Assisted
[50,75,80,87,88]
Viscosity of the Bioink
30–6 × 107 mPa•s <10 mPa•s 1–300 mPa•s
Cell Density High, cell spheroids Low, <106 cells/ml Medium (108 cells/mL)
Resolution 200–103 µm 10–50 µm 10–100 µm
Speed of Fabrication 10–700 mm/s 105 droplets/s 200–1600 mm/s
Cell Viability 80–90% >85% >95%
Price Moderate Low High
Advantages High-
viscosity printing, print high
cell densities
Inexpensive, high printing
speed, moderate cell viability
High printing speed and
precision, high cell viability
Figure 3.
A representation of laser-assisted bioprinting with bioink droplets deposited by laser pulses
onto a collector slide [38].
2.3.1. Laser-Assisted Bioprinting of Adipose-Derived Stem Cells
Koch et al. determined that laser-assisted bioprinting did not initiate differentiation
due to conservation of the hADSC immunophenotypes CD44, CD105, CD29, and CD90 [
59
].
The viability of hADSCs post printing was determined to be 99.7% in one study conducted.
Furthermore, the proliferative ability of the stem cells was shown to be unimpacted. DNA
damage in printed hADSCs was not significant relative to the control cells showing an
absence of genotoxicity indicating laser exposure may not have adverse effects [
60
]. A
study fabricating corneal tissue with hADSCs in human Col I hydrogels showed high
viability immediately after printing with a Nd:YAG and Er:YAG laser. The hADSCs
retained proliferative capabilities as Ki67 was expressed on day 1 and day 4 post printing.
The proliferation rate significantly increased after 4 days of culturing which may be due to
the biocompatible nature of collagen [
61
,
62
]. A separate study conducted by Gruene et al.
laser printed hADSCs in alginate/EDTA blood plasma hydrogels to evaluate the effects of
laser printing. There was no change in cell behavior, which was determined by measuring
cell proliferation, which showed no significant difference in the laser printed cells. The
cells survived the stress of the laser printing and retained their differentiation potential
into adipocytes, which was verified by Oil red O staining and RT-qPCR for adipogenic
genes [63].
Overall, laser printing hADSCs has no detrimental effect on their proliferation, via-
bility, and differentiation, making it an optimal printing method for creating cell-laden
scaffolds. However, there may be unwanted differentiation due to the physical forces
Sensors 2021,21, 7477 8 of 20
present during the printing process [
64
]. More studies should be conducted to determine
the effects of laser exposure in the manner of osteogenic differentiation of ADSCs.
2.3.2. Laser-Assisted Bioprinting of Bone-Marrow-Derived Stem Cells
Gruene et al. printed porcine BMSC (pBMSC) hydrogel scaffolds through laser-
assisted bioprinting and determined cell viability and differentiation potential post printing.
Cell viability and proliferation exhibited no significant difference, and no changes were
detected in pBMSC phenotype. The pBMSCs displayed an increase in aggrecan expression
with a lack of collagen type II expression. The findings indicated that MSCs in a scaffold
are predisposed to shift to chondrogenic differentiation in a 3D culture. Additionally, it
was determined that laser bioprinting caused no significant spontaneous differentiation
into osteoblasts by measuring ALP activity [
65
]. A separate study conducted by Koch
et al. determined that there was no significant difference in apoptosis, proliferation, and
genotoxicity in hBMSCs post printing with a Nd:YAG-laser. The hBMSCs demonstrated
a survival rate of 90% after printing [
59
]. Laser bioprinting confers printing with high
resolution and precision, as shown by a printing resolution of 138
µ
m and precision of
16
µ
m in one study with BMSCs [
66
]. Ali et al. used slow jet conditions, which are more
stable, to minimize droplet impact energy with mice BMSCs (mBMSC). The slow jetting
conditions have decreased laser pulse energy, which reduces shear stress. The mBMSCs
were printed with high cell viability, which was measured 24 h after printing, and possessed
high resolution [
67
]. Laser bioprinting can be used to deposit BMSCs directly
in vivo
to
enhance osteogenesis. Keriquel et al. devised a method to print nano-Hydroxyapatite
(nHA) layers directly onto a mouse critical sized calvaria defect. The experimentation
demonstrated laser exposure to the dura mater caused temporary inflammation and no
permanent tissue damage in mouse brain [
68
]. This was further expanded by printing
BMSCs in situ in a ring or disk geometry to induce osteogenesis
in vivo
. The in situ printed
BMSC nHA disks showed significant osteogenesis than the ring shaped BMSC nHA. It
is hypothesized that due to the disk cell homogeny and proximity, the BMSCs secreted
paracrine factors to induce osteogenic differentiation [
69
]. This novel technique should be
explored in greater depth with different biomaterials and BMSCs to gauge its full potential.
A summary of each bioprinting technique and its effects on ADSCs and BMSCs is provided
in Table 2.
Table 1. Bioprinting techniques.
Extrusion [50,70–80] Inkjet [50,51,80–86]Laser Assisted
[50,75,80,87,88]
Viscosity of the
Bioink 30–6 ×107mPa·s<10 mPa·s 1–300 mPa·s
Cell Density High, cell spheroids Low, <106cells/ml
Medium (10
8
cells/mL)
Resolution 200–103µm10–50 µm 10–100 µm
Speed of
Fabrication 10–700 mm/s 105droplets/s 200–1600 mm/s
Cell Viability 80–90% >85% >95%
Price Moderate Low High
Advantages
High-viscosity
printing, print high
cell densities
Inexpensive, high
printing speed,
moderate cell
viability
High printing speed
and precision, high cell
viability
Disadvantages
High shear stress,
lower cell viability,
slow printing
Low cell density, low
viscosity biomaterials,
nozzle clogging
Expensive, complex
laser control
Sensors 2021,21, 7477 9 of 20
Table 2. Effects of bioprinting techniques on adipose and bone marrow stem cells.
Bioprinting Method
Extrusion Inkjet Laser References
Adipose Stem Cells
Drop in viability due to
shear stress, cells can
attach to hydrogels
normally and grow,
printed monolayers
show a higher cell
viability, retention of
differentiation ability
Favorable cell adhesion
depends on the
biomaterial, increase in
cell proliferation after
24 h, may create
incomplete constructs
due to printing lower
cell densities
Does not initiate
differentiation, no
effect on proliferation,
no significant DNA
damage
[40–44,51–53,59–62]
Bone Marrow Stem
Cells
Sheer stress may
encourage cells into
osteoblast lineage,
long-term
differentiation potential
is retained, lower cell
viability, cell
proliferation increases
within 28 days
Cell proliferation and
viability affected by
higher pressures,
medium shear pressure
encourages
differentiation,
unchanged stem cell
phenotype post
printing, osteogenic
differentiation not
affected by printing
No changes in
phenotype, no
significant effect on cell
proliferation, high cell
viability, no significant
genotoxicity or
apoptosis occurred
[37,45–48,55,56,59,65–67]
3. Scaffolds
Prior to 3D bioprinting, a bioink must be created. Bioink is referred to as the mixture
of cells and biopolymers or biomaterials. The properties of the bioink vary and are largely
dependent on the ink’s constituents. Synthetic or natural materials may be used to create
synthetic or natural bioink. Once the bioink has undergone the bioprinting process, a
scaffold is formed. Scaffolds are the structural form of a bioink and provide micro and
macro-architecture for cell attachment
in vitro
and for
in vivo
implantation. In this paper,
we define “scaffolds” as multipurpose with the function of being tested
in vitro
or
in vivo
.
The following sections evaluate the use of scaffolds with varying bioink constituents in
bone tissue regeneration.
3.1. Adipose-Derived Stem Cells
3.1.1. Natural Scaffolds
Alginate
Jia et al. bioprinted four RGD-alginate hydrogels with varying oxidation and concen-
trations with hADSCs in a lattice structure through the use of a custom inkjet printer. The
alginate was oxidized with the addition of sodium periodate at room temperature. How-
ever, crosslinking was performed with the addition of a 100 mM CaCl
2
gelatin substrate.
The substitution of sodium ions for calcium ions allows the strengthening of the polymer
yielding a solid hydrogel. Post cross-linking, the rigid form of the alginate hydrogel al-
lows it to resist compressive forces. Alginate with 5% oxidation and 15% concentration
had the greatest hADSC proliferation and spreading compared to the 0% oxidized and
8% concentration alginate scaffold. This was potentially due to the greater porosity and
degradation rate. The differences in scaffolds impacted cell morphology and behavior.
The 0%-ox.–8%-conc.
scaffold conferred a round morphology associated with chondrogen-
esis. However, hADSCs with a larger spreading area as seen in the 5%-ox.–15%-conc. group
were correlated with osteogenesis [
89
,
90
]. Kim et al. characterized porcine ADSC morphol-
ogy and transcriptome in a 2% alginate scaffold that was reconstituted from lyophilized
alginate. The pADSCs formed osteogenic nodules and displayed increased expression
of ALP by day 4 after exposure to osteogenic media. An early increase in the expression
of BGLAP, COL1A1, SPARC, and SPP1 was noticeable by day 2. Cell morphology was
Sensors 2021,21, 7477 10 of 20
rounded in the 3D alginate scaffold contrary to fibroblast-like shape in 2D culture prior to
osteogenesis. Alginate stiffness and environment play a role in cell attachment and osteo-
genesis, since a stiff environment may encourage cells to differentiate into an osteoblast
lineage and limit migration [
91
,
92
]. Guneta et al. show hADSCs prefer osteogenic differen-
tiation in alginate scaffolds with a stiffness greater than 11.61 kPa [
93
]. A study conducted
by Ghiasi et al. determined the effects of multiple scaffolds, including alginate, on hADSC
viability and proliferation. The alginate scaffold was reconstituted by creating an alginate
mixture from powder and cells. After 14 days of culturing in a 1.2% w/walginate scaffold,
hADSCs expressed the pluripotent markers OCT4A, NANOG, and SOX2, indicating the
preservation of stemness. Compared to the FG, PLGA, and active and inactive PRP scaf-
folds, the cell-laden alginate gel displayed the lowest viability and proliferation, which
may be due to the lack of integrin binding sites for cells [
94
,
95
]. Despite alginate being a
commonly used material in tissue engineering due to its biocompatibility and low toxicity,
Arg-Gly Asp (RGD) conjugation may often be required for increased cell adhesion [96].
Collagen
Alkaline phosphatase can be cross-linked onto collagen fibers to improve osteogenic
differentiation [
97
]. A study conducted by Jafary et al. evaluated hADSC differentiation
on collagen fiber scaffolds with covalently immobilized alkaline phosphatase. Greater
osteocalcin, ALP, and collagen expression were detected on the third day with expression
decreasing on the seventh day. The levels of RUNX2 were downregulated compared to
the collagen only group on the third day with no significant difference on the seventh
day. Expression of TNF-
α
, an inhibitor gene for osteogenesis, decreased on the third
day. Altogether, osteogenesis was improved when ALP was immobilized on collagen
scaffolds, with osteogenic gene upregulation and inhibitor gene downregulation indicating
the potential of seeding enzymes on cell-laden collagen scaffolds [
98
]. Geometrically
honeycomb collagen scaffolds seeded with hADSCs can induce osteogenesis. Greater Cbfa-
1 expression was present in the cells seeded in the collagen scaffold indicating osteogenesis
activation. Cell migration was enhanced, as by day 14, the hADSCs were migrating
deeper into the collagen scaffold. Additionally,
in vivo
findings showed positive von Kossa
staining and osteocalcin immunostaining indicating the formation of mineralized nodules
and bone formation [
99
]. Furthermore, aligned collagen type I fibers placed in synthetic
electrospun PLGA/PCL scaffolds can have effects on cell morphology and migration
which has implications on guided bone regeneration. Rat ADSCs had an oval nucleus with
directionally oriented bipolar morphology in comparison to the multipolar morphology
present in non-aligned fibers. The presence of the collagen fibers increased the expression
of osteogenic genes; however aligned collagen fibers were better than the random fibers in
osteogenesis [100].
Gelatin
Canine ADSCs (cADSC) were used in making gelatin-induced osteogenic cell sheets to
observe osteogenic differentiation and proliferation. The addition of gelatin to osteogenic
medium results in greater cell proliferation, upregulation of BMP-7, and positive alizarin
red staining [
101
]. A similar study used frozen–thawed gelatin-induced osteogenic cell
sheets with cADSCs for
in vivo
repair in a canine model. Large callus formation was
present in all groups, but better cortical bone connectivity was present in the gelatin
treated groups relative to the untreated ADSCs. These findings indicate the potential of
gelatin in fracture repair due to greater ossification and mature bone presence
in vivo
[
102
].
Furthermore, Wofford et al. evaluated commercially acquired purified gelatin scaffolds
seeded with hADSCs in a rat maxillary alveolar bone defect model. The gelatin scaffold was
not cytotoxic and encouraged hADSC growth and attachment post seeding. Expression
of OPN and RUNX2 were significantly upregulated with osteogenesis occurring with
no adverse effects of the gelatin scaffold. Rats treated with hADSC + Gelatin scaffolds
had greater bone formation early on with Masson trichrome staining verifying organized
Sensors 2021,21, 7477 11 of 20
ossification [
103
]. A recent study conducted by Wang et al. tested the effectiveness of
reconstituted gelatin scaffolds with hADSCs. The scaffold was biocompatible, as cell
adhesion and proliferation were sustained. Interestingly, OCT4, nestin, and SOX9 were
downregulated, while osteocalcin was upregulated more than 2-fold in a 14-day period. The
findings suggest gelatin scaffolds emphasize osteogenic differentiation compared to other
lineages. Von Kossa and alizarin red staining detected the formation of calcium nodules
within 14 days. The scaffold was implanted in a rat calvarial defect and greater bone
formation was present relative to the untreated group indicating enhanced osteogenesis
with the gelatin scaffold [104].
Hyaluronic Acid
Aguiari et al. seeded hADSCs onto a hyaluronan-based sponge and observed differ-
entiation into osteogenic, chondrogenic, and adipogenic lineages. In the osteogenic culture,
positive ALP activity was detected along with osteonectin, osteopontin, and osteocalcin
expression. Markers for adipocyte and chondrocyte differentiation were not. Mineralized
bone matrix formation was surrounding the cells indicating osteogenesis in the hyaluro-
nan scaffold [
105
]. mADSCs in hyaluronan-coated or hyaluronan-supplemented cultures
demonstrated greater osteogenic potential along with long term cell preservation. Cell
senescence was significantly less in both hyaluronan groups, indicating that hyaluro-
nan suppresses cell senescence for long-term proliferation. Greater calcium deposition
was shown in both hyaluronan groups, detected by silver nitrate staining. In addition,
osteogenic potential was preserved in passages greater than 5 [
106
]. Although this ex-
periment was not conducted in a 3D culture, it highlights the potential application of
hyaluronan in preserving cell osteogenic differentiation and lifespan for future prospects
with ADSCs. Similarly, hADSCs were cultured in hyaluronic acid-derived scaffolds for
corneal repair. The ADSCs cultured on the commercially acquired Hystem-HP scaffold
had the greatest survivability since the environment promoted cell growth. Cells in the
hydrogel scaffold survived long-term up to 10 weeks, indicating the preservative effects
of hyaluronan [
107
]. More experimentation with ADSCs in hyaluronan-based scaffolds
should be conducted to further characterize its uses in bone regeneration and osteoblast
formation.
3.1.2. Natural Composite Scaffolds
Alginate/Collagen
A study conducted by Yeo et al. fabricated hydrogels with a collagen core seeded
with hADSCs with an outer alginate sheath through a custom-made 3D bioprinter. hAD-
SCs in the collagen center were healthy and proliferation increased over a 7-day period
significantly compared to alginate-based control scaffolds. Cell viability of hADSCs in
the composite was at 91% compared to the 83% in the alginate hADSC control scaffold,
showing collagen’s cell-supportive nature. Live/dead fluorescent images of the hADSCs in
the composite scaffold showed greater live cells on day 7. In addition, hepatogenic genes
ALB and TDO2 were higher in the composite scaffolds indicating potential hepatogenic
differentiation. Currently, this may have future use in liver regeneration studies [
108
].
For bone regeneration, osteogenic genes should be evaluated along with the impact of
alginate/collagen concentration on hADSC phenotype in accordance with osteoblast differ-
entiation.
Collagen/Hyaluronic Acid
Xu et al. used ADSCs in reconstituted collagen/hyaluronic acid composites for
vocal fold regeneration. ADSC viability a week post-seeding was improved along with
greater proliferation which suggests the scaffolds foster ADSC growth [
109
]. Another
study conducted by Amann et al. co-cultured hADSCs with human articular chondrocytes
in collagen/hyaluronic acid scaffolds reconstituted via gelation. The greater hyaluronic
acid concentration was associated with lower SOX9 expression indicating chondrogenesis
Sensors 2021,21, 7477 12 of 20
inhibition. Metalloproteinase 13 expression increased which is related to matrix remodeling.
Lower concentrations of hyaluronan, such as 1%, resulted in SOX9 upregulation and
glycosaminoglycan production which is associated with chondrogenesis. In addition, the
low O
2
levels detected in the scaffolds may contribute to improved chondrogenesis. The
co-culturing of articular chondrocytes and hADSCs prefer chondrogenesis, but hyaluronan
concentration may be modified to prevent chondrogenesis [
110
]. Hyaluronan concentration
in a HA/collagen composite scaffold and osteoblast co-culturing with hADSCs should be
investigated for use in bone regeneration. The effects of each biomaterial on ADSCs are
summarized in Table 3.
Table 3. Adipose stem cells in scaffolds.
Biomaterial Effects Bioprinting Method * References
Alginate
Increased BGLAP, COL1A1, SPP1 expression, round
morphology, stiffness may encourage osteoblast
differentiation, stemness preservation, low viability
Inkjet [91,93,94]
Collagen
Osteogenic genes are upregulated with inhibition of
TNF-α, enhanced in vivo bone regeneration,
multipolar or bipolar morphology
- [98–100]
Gelatin
Good cell proliferation and adhesion, ossification
in vivo, enhance osteogenesis, early upregulation of
OPN and RUNX2
- [101–104]
Hyaluronic Acid
Can cause osteogenic, chondrogenic, or adipogenic
differentiation, mineralized matrix formation and
calcium deposition, long term cell preservation
- [105–107]
Alginate/Collagen High viability and cell proliferation Custom [108]
Collagen/Hyaluronic Acid
Strong cell proliferation, associated with
chondrogenesis, greater hyaluronic concentration
inhibits chondrogenesis
- [109,110]
* If applicable.
3.2. Bone-Marrow-Derived Stem Cells
3.2.1. Natural Scaffolds
Alginate
pBMSCs cultured on a 2% alginate scaffold displayed increased expression of os-
teogenic genes but had a delayed increase in ALP and BGLAP. The pBMSCs formed
osteogenic nodules by day 4 and stained positive for ALP by day 14. ALP presence was
detected a few days after the presence of ALP in a 2D culture, indicating the possibility
of poor diffusion of differentiation factors or dyes through the alginate hydrogel. Cell
morphology was spherical prior to osteoblast differentiation in the 3D culture. pBMSCs
exhibited substantial viability in the alginate under osteogenic conditions over the period
of differentiation as detected by an MTT assay [
91
]. A study comparing alginate-CaCl
2
,
alginate-CaSO
4
, alginate-gelatin, and alginate-nanocellulose seeded with BMSCs was con-
ducted to determine osteogenic and grafting potential through extrusion 3D bioprinting.
There were fewer viable cells in the alginate-CaCl
2
and alginate-gel scaffold, but these
groups had the lowest apoptotic levels. The greatest cell death occurred in the alg-CaSO
4
and alg-nanocellulose scaffolds after 7 days possibly due to the mechanical properties of the
hydrogel limiting nutrient exchange. The alg-CaCl
2
scaffold displayed the greatest calcium
deposition and positive osteocalcin expression indicating it is favorable for osteogenesis.
To further evaluate the scaffolds, a scaphoid bone was bioprinted using the alg-CaCl
2
bioink with BMSCs with viable cells post printing. After 2 weeks, calcification was present
with positive alizarin red staining. Despite the increase in calcification, deposition was
not homogenous and was focused near the peripheral edges [
111
]. Future applications
of cell-laden bone fabrication should be explored with a focus on vascularization and
Sensors 2021,21, 7477 13 of 20
improved mechanical properties as alginate concentration may impact nutrient and waste
diffusion [112].
Collagen
Collagen is rich in integrin binding sites, which promotes cell adhesion and can be
combined with other materials to offer improved mechanical and biological properties [
113
].
The addition of type I collagen to a culture can induce osteoblast differentiation in bone
marrow stromal cells. This was indicated by upregulation of differentiation markers ALP,
Cbfa-1, OCN, and mineralization that was present at three weeks [
114
]. A study by George,
Kuboki, and Miyata created lyophilized honeycomb collagen scaffolds with rBMSCs and
determined the differentiation potential into osteoblasts. Greater collagen content and ALP
activity was detected on the honeycomb collagen scaffold along with calcium-deficient
hydroxyapatite synthesis. The scaffolds contained pore sizes of 200–400
µ
m and promoted
osteogenesis as verified through X-ray diffraction and Von Kossa staining [
115
]. Apart
from type I collagen, type II collagen-coated culture plates with hBMSCs demonstrated
significant RUNX2 upregulation along with osteocalcin by day 6, indicating that type II
collagen affects calcium deposition. The hBMSCs also expressed upregulation of the type
II collagen receptor integrin
α
2
β
1 by day 4 during osteogenic differentiation indicating
attachment to the type II collagen. In a powder reconstituted HA/TCP scaffold coated with
type II collagen, increased mineral deposition and cell calcification were detected relative
to the control or type I collagen coated scaffolds. Additionally, denser bone formation
indicating mature bone was present during the assessment of segmental bone defect repair
in rats. On day 7, there was a significant improvement in walking function for mice treated
with the type II collagen-coated HA/TCP and type I collagen-coated HA/TCP scaffold.
However, sciatic function index scores were consistently higher for the type II collagen
coated scaffold but after day 28 equivalent outcomes were shown in walking function [
116
].
The use of collagen fibers and BMSCs in biomimetic scaffolds is promising with future
applications expanding on bone reconstruction.
Gelatin
Mazaki et al. created a lyophilized gelatin-furfurylamine hydrogel photo-crosslinked
with Rose Bengal. The hydrogels were seeded with BMSCs, BMP4, and collagen binding
domains. Rose Bengal alone is cytotoxic to cells, but the addition of the gelatin-FA hydrogel
confers cytoprotecting function as cell viability increased significantly. The scaffolds were
implanted into a rabbit osteochondral defect and after 12 weeks the BMSC-laden gelatin-FA
hydrogel exhibited cartilage and bone growth similar to the surrounding areas, indicating
osteogenesis and chondrogenesis [
117
]. Another study by Yao et al. synthesized a BMSC-
laden lyophilized gelatin hydrogel with horseradish peroxidase and galactose oxidase
enzymatic crosslinking in dermal regeneration. The encapsulated BMSCs had greater
proliferation and viability which may be due to the less exposure of H
2
O
2
. There was a
faster healing time in dermal wounds, but the implications for bone regeneration should be
explored, as the presence of reactive oxygen species increases during a fracture [118,119].
Hyaluronic Acid
Hyaluronic acid in a 3D culture can have positive implications in inducing osteoge-
nesis in BMSCs. Hyaluronan hydrogels synthesized from powder and conjugated with
N-cadherin mimetic peptides demonstrate an increase in osteogenic markers
in vitro
and
mineralization both
in vitro
and
in vivo
. There is significant upregulation of type I collagen,
osteocalcin, ALP, and RUNX2 after 4 days of osteogenic differentiation in the Cad + RGD
hyaluronan scaffold. Cell viability is markedly increased due to increased cell adhesion in
the RGD peptide conjugated HA scaffold. There is greater mineralization and angiogenesis
occurring in the Cad + RGD group, indicating successful bone matrix formation. Micro
CT skull reconstruction in rat calvarial defects displayed greater bone formation related to
the Cad + RGD group, showing
in vivo
efficacy in critical size defect repair. Expression of
Sensors 2021,21, 7477 14 of 20
osteocalcin and RUNX2, blood vessel formation, and higher bone volume in the Cad + RGD
HA group
in vivo
signifies increased bone matrix deposition, along with the formation of
osteoblasts [
120
]. A separate study conducted by Cavallo et al. evaluated the use of bone
marrow concentrate (BMC) containing stem cells, immune cells, and resident marrow cells
in HA scaffolds. Hyaff-11 scaffolds were seeded with the BMC replicating the bone mar-
row microenvironment to induce osteogenesis to repair osteochondral lesions. The BMC
cells were viable post-seeding and proliferated successfully until day 40. Mineralization
occurred after day 40 with the presence of dark nodules confirmed via Von Kossa staining.
Type I collagen, BSP, and ALP levels increased around the 40-day mark. RUNX2 levels
were high at day 0 but decreased until day 52 indicating early bone formation [
121
]. This
indicates hyaluronan has positive implications for early-onset osteogenesis.
3.2.2. Natural Composite Scaffolds
Alginate/Collagen
Perez et al. designed a hydrogel carrier with a collagen–BMSC core and an alginate
outer shell for cell delivery. The encapsulated BMSCs had a cell viability of 70% throughout
a 21-day period; however, proliferation was persistent for up to 21 days. Cell survival in
the carriers depended on the alginate/collagen concentrations, as alginate concentrations
that are too high result in hydrogel stiffness and nutrient diffusion limits. There was
greater BSP, OPN, and OCN expression in the hydrogel composite carriers compared to the
collagen gel by day 14, indicating a greater initial propensity for osteogenesis. The
in vivo
rat calvarial model had the greatest osteogenesis in the BMSC-encapsulated carrier group
which was differentiated for 7 days. The nondifferentiated BMSC carrier group displayed
a 25% increase in bone volume but was exceeded by the differentiated carrier group with
up to 45% ossification [
122
]. In conclusion, more experimentation should be conducted on
BMSCs in alginate/collagen scaffolds to determine the full effects on osteogenesis
in vitro
and in vivo.
Collagen/Hyaluronic Acid
rBMSCs in collagen/hyaluronic acid scaffolds fabricated by freeze-drying demon-
strate early chondrogenesis and drastic upregulation of SOX9 and collagen type II. Cell
proliferation increased within 28 days in the collagen/hyaluronan scaffold compared to
the collagen-only scaffold. In addition, greater cell infiltration was present in the compos-
ite scaffold despite smaller pore size indicating an influence on cellular migration [
123
].
Additionally, the findings of Zhang et al. demonstrate that the use of collagen/hyaluronic
acid seeded with BMSCs are suitable for cartilage tissue engineering [
124
]. Differentiation
of the BMSCs into chondrocytes makes this scaffold ideal for chondrogenesis compared to
osteogenesis. The use of strict BMSC-laden collagen/hyaluronic acid hydrogels in bone
tissue engineering is scarce and should be further evaluated for bone defect repair [
125
].
Table 4summarizes the effects of each biomaterial on BMSCs.
Table 4. Bone marrow stem cells in scaffolds.
Biomaterial Effects Bioprinting Method * References
Alginate
Early osteogenic nodule formation, poor diffusion of
differentiation factors/dyes, spherical morphology,
lower cell viability, presence of calcium deposition
Extrusion [91,111,112]
Collagen
Can induce osteogenic differentiation, mineralization
present by 21 days, RUNX2 upregulation, cells bind to
integrin proteins
- [114–116]
Gelatin Cytoprotective function, osteogenesis and
chondrogenesis, strong proliferation and cell viability - [117,118]
Hyaluronic Acid Increase in osteogenic markers, blood vessel formation
and mineralization in vivo, early bone formation - [120,121]
Sensors 2021,21, 7477 15 of 20
Table 4. Cont.
Biomaterial Effects Bioprinting Method * References
Alginate/Collagen
Lower cell viability but persistent cell proliferation,
survival depends on alginate/collagen concentrations,
greater BSP, OCN, and OPN expression
- [122]
Collagen/Hyaluronic
Acid
Early chondrogenesis, upregulation of SOX9 and
collagen type II, greater cell migration - [123]
* If applicable.
4. Conclusions
Current bioprinting methods are capable of printing complex 3D structures seeded
with stem cells for bone tissue engineering; however, they are yet to surpass autologous
bone grafts. Extrusion, inkjet, and laser-assisted bioprinting each have their advantages
and disadvantages in their use for printing stem cells. Furthermore, the implementation
of nanotechnology and functionalized materials are opening new avenues with bioprint-
ing [
126
,
127
]. Their implications on cell viability and differentiation should be considered
prior to the use of these methods. Mesenchymal stem cells such as adipose-derived and
bone-marrow-derived stem cells provide routes for enhancing osteogenesis due to their
differentiation ability. When these stem cells are placed into a scaffold, osteogenesis can be
induced and cause fracture reparation. BMSCs and ADSCs exhibit similar and different
properties in the various scaffolds discussed. Both gravitate towards chondrogenesis in
collagen/hyaluronic acid composites and have strong proliferation in alginate/collagen
composites and gelatin. Interestingly, BMSCs have the potential to differentiate into a chon-
drogenic line, as well as an osteogenic one in gelatin, while the studies discussing ADSCs
involve enhanced osteogenesis. Overall, both cell types have upregulated osteogenic genes,
such as RUNX2, OPN, and OCN, contributing to osteogenesis. Aside from osteogenesis,
chondrogenesis may occur, depending on the biomaterials used. Natural polymers can
be printed individually or mixed to form composites with cells. These natural scaffolds
can enhance differentiation, cell viability, and cell adhesion depending on the polymer’s
properties. The biodegradability, biocompatibility, and osteoinductive effects of the de-
sired scaffold should be evaluated prior to use with the stem cells in order to maximize
osteogenic potential. In addition, composite scaffolds provide a mix of properties from
both biomaterials which can confer better traits than an individual biomaterial. Therefore,
the printing method, combination of biomaterials, and type of stem cells must be carefully
considered prior to osteogenic differentiation for fracture repair. In summary, while many
natural polymers have been used to evaluate BMSCs and ADSCs
in vitro
and
in vivo
, more
experimentation must be performed to maximize the efficiency of using these cells in
biomaterials. Furthermore, the process of fracture repair is dynamic, and investigation
should be focused on the interaction between the biomaterials, cells, and the underlying
bone tissue in
in vivo
models. This allows insight into the molecular mechanisms of repair
mediated by the biomaterials and cells. Currently, the interactions between 3D bioprinting
and cell-laden bioink have been characterized across extrusion, inkjet, and laser-assisted
bioprinting with respect to viability, genotoxicity, and migration. However, more studies
need to be conducted to further evaluate the impact on osteogenesis between cell-laden
biomaterials and 3D bioprinting, as many scaffold compositions need further testing. Be-
tween these three methods, careful consideration must be taken when determining the
composition of the bioink to generate a scaffold. Only by optimizing this process can
experiments create an ideal resulting scaffold to maximize bone regeneration.
Author Contributions:
All authors contributed equally to the manuscript. Conceptualization, A.I.;
writing—original draft preparation, S.T.; writing—review and editing, S.K. and A.I.; supervision, A.I.
All authors have read and agreed to the published version of the manuscript.
Funding: This research received no external funding.
Sensors 2021,21, 7477 16 of 20
Institutional Review Board Statement: Not applicable.
Informed Consent Statement: Not Applicable.
Data Availability Statement: Not Applicable.
Conflicts of Interest: The authors declare no conflict of interest.
References
1.
Burge, R.; Dawson-Hughes, B.; Solomon, D.H.; Wong, J.B.; King, A.; Tosteson, A. Incidence and economic burden of osteoporosis-
related fractures in the United States, 2005–2025. J. Bone Min. Res. 2007,22, 465–475. [CrossRef]
2. Myeroff, C.; Archdeacon, M. Autogenous bone graft: Donor sites and techniques. JBJS 2011,93, 2227–2236. [CrossRef]
3.
Huang, Y.-C.; Chen, C.-Y.; Lin, K.-C.; Renn, J.-H.; Tarng, Y.-W.; Hsu, C.-J.; Chang, W.-N.; Yang, S.-W. Comparing morbidities of
bone graft harvesting from the anterior iliac crest and proximal tibia: A retrospective study. J. Orthop. Surg. Res.
2018
,13, 115.
[CrossRef]
4.
Salawu, O.; Babalola, O.; Ahmed, B.; Ibraheem, G.; Kadir, D. Comparative study of proximal tibia and iliac crest bone graft donor
sites in treatment of orthopaedic pathologies. Malays. Orthop. J. 2017,11, 15. [PubMed]
5.
Carlisle, E.R.; Fischrund, J.S. Bone Graft and Fusion Enhancement. In Surgical Management of Spinal Deformities; Elsevier:
Amsterdam, The Netherlands, 2009; pp. 433–448.
6.
Boone, D.W. Complications of iliac crest graft and bone grafting alternatives in foot and ankle surgery. Foot Ankle Clin.
2003
,8,
1–14. [CrossRef]
7.
Zhang, X.; Awad, H.A.; O’Keefe, R.J.; Guldberg, R.E.; Schwarz, E.M. A perspective: Engineering periosteum for structural bone
graft healing. Clin. Orthop. Relat. Res. 2008,466, 1777–1787. [CrossRef] [PubMed]
8.
Oryan, A.; Alidadi, S.; Moshiri, A.; Maffulli, N. Bone regenerative medicine: Classic options, novel strategies, and future
directions. J. Orthop. Surg. Res. 2014,9, 18. [CrossRef]
9.
Fernandez de Grado, G.; Keller, L.; Idoux-Gillet, Y.; Wagner, Q.; Musset, A.-M.; Benkirane-Jessel, N.; Bornert, F.; Offner, D. Bone
substitutes: A review of their characteristics, clinical use, and perspectives for large bone defects management. J. Tissue Eng.
2018
,
9, 2041731418776819. [CrossRef] [PubMed]
10.
Moroni, L.; Boland, T.; Burdick, J.A.; De Maria, C.; Derby, B.; Forgacs, G.; Groll, J.; Li, Q.; Malda, J.; Mironov, V.A. Biofabrication:
A guide to technology and terminology. Trends Biotechnol. 2018,36, 384–402. [CrossRef]
11. Murphy, S.V.; Atala, A. 3D bioprinting of tissues and organs. Nat. Biotechnol. 2014,32, 773–785. [CrossRef] [PubMed]
12.
Takizawa, T.; Nakayama, N.; Haniu, H.; Aoki, K.; Okamoto, M.; Nomura, H.; Tanaka, M.; Sobajima, A.; Yoshida, K.; Kamanaka, T.
Titanium fiber plates for bone tissue repair. Adv. Mater. 2018,30, 1703608. [CrossRef] [PubMed]
13.
Polo-Corrales, L.; Latorre-Esteves, M.; Ramirez-Vick, J.E. Scaffold design for bone regeneration. J. Nanosci. Nanotechnol.
2014
,14,
15–56. [CrossRef] [PubMed]
14.
Ouasti, S.; Donno, R.; Cellesi, F.; Sherratt, M.J.; Terenghi, G.; Tirelli, N. Network connectivity, mechanical properties and cell
adhesion for hyaluronic acid/PEG hydrogels. Biomaterials 2011,32, 6456–6470. [CrossRef] [PubMed]
15.
Moxon, S.R.; Corbett, N.J.; Fisher, K.; Potjewyd, G.; Domingos, M.; Hooper, N.M. Blended alginate/collagen hydrogels promote
neurogenesis and neuronal maturation. Mater. Sci. Eng. C 2019,104, 109904. [CrossRef] [PubMed]
16.
Rowley, J.A.; Mooney, D.J. Alginate type and RGD density control myoblast phenotype. J. Biomed. Mater. Res.
2002
,60, 217–223.
[CrossRef] [PubMed]
17. Chocholata, P.; Kulda, V.; Babuska, V. Fabrication of scaffolds for bone-tissue regeneration. Materials 2019,12, 568. [CrossRef]
18.
Donnaloja, F.; Jacchetti, E.; Soncini, M.; Raimondi, M.T. Natural and Synthetic Polymers for Bone Scaffolds Optimization. Polymers
2020,12, 905. [CrossRef]
19.
Qu, H.; Fu, H.; Han, Z.; Sun, Y. Biomaterials for bone tissue engineering scaffolds: A review. RSC Adv.
2019
,9, 26252–26262.
[CrossRef]
20. Davis, H.; Leach, J. Hybrid and composite biomaterials in tissue engineering. Top. Multifunct. Biomater. Devices 2008,10, 1–26.
21.
Nebhani, L.; Choudhary, V.; Adler, H.-J.P.; Kuckling, D. pH-and Metal Ion-Sensitive Hydrogels based on N-[2-(dimethylaminoethyl)
acrylamide]. Polymers 2016,8, 233. [CrossRef] [PubMed]
22.
Xing, Q.; Yates, K.; Vogt, C.; Qian, Z.; Frost, M.C.; Zhao, F. Increasing mechanical strength of gelatin hydrogels by divalent metal
ion removal. Sci. Rep. 2014,4, 4706. [CrossRef] [PubMed]
23.
Kamal, M.M.; Kassem, D.H. Therapeutic potential of wharton’s jelly mesenchymal stem cells for diabetes: Achievements and
challenges. Front. Cell Dev. Biol. 2020,8, 16. [CrossRef]
24.
Ullah, I.; Subbarao, R.B.; Rho, G.J. Human mesenchymal stem cells-current trends and future prospective. Biosci. Rep.
2015
,35,
e00191. [CrossRef] [PubMed]
25.
Li, X.; Bai, J.; Ji, X.; Li, R.; Xuan, Y.; Wang, Y. Comprehensive characterization of four different populations of human mesenchymal
stem cells as regards their immune properties, proliferation and differentiation. Int. J. Mol. Med.
2014
,34, 695–704. [CrossRef]
[PubMed]
26.
Yousefi, A.-M.; James, P.F.; Akbarzadeh, R.; Subramanian, A.; Flavin, C.; Oudadesse, H. Prospect of stem cells in bone tissue
engineering: A review. Stem Cells Int. 2016,2016, 6180487. [CrossRef]
Sensors 2021,21, 7477 17 of 20
27.
Yu, L.; Wu, Y.; Liu, J.; Li, B.; Ma, B.; Li, Y.; Huang, Z.; He, Y.; Wang, H.; Wu, Z. 3D culture of bone marrow-derived mesenchymal
stem cells (BMSCs) could improve bone regeneration in 3D-printed porous Ti6Al4V scaffolds. Stem Cells Int.
2018
,2018, 2074021.
[CrossRef] [PubMed]
28.
Olivares, A.L.; Lacroix, D. Simulation of cell seeding within a three-dimensional porous scaffold: A fluid-particle analysis. Tissue
Eng. Part C Methods 2012,18, 624–631. [CrossRef] [PubMed]
29.
Derakhshanfar, S.; Mbeleck, R.; Xu, K.; Zhang, X.; Zhong, W.; Xing, M. 3D bioprinting for biomedical devices and tissue
engineering: A review of recent trends and advances. Bioact. Mater. 2018,3, 144–156. [CrossRef]
30.
Chimene, D.; Kaunas, R.; Gaharwar, A.K. Hydrogel bioink reinforcement for additive manufacturing: A focused review of
emerging strategies. Adv. Mater. 2020,32, 1902026. [CrossRef]
31.
Zhu, K.; Shin, S.R.; van Kempen, T.; Li, Y.C.; Ponraj, V.; Nasajpour, A.; Mandla, S.; Hu, N.; Liu, X.; Leijten, J. Gold nanocomposite
bioink for printing 3D cardiac constructs. Adv. Funct. Mater. 2017,27, 1605352. [CrossRef] [PubMed]
32.
Mannoor, M.S.; Jiang, Z.; James, T.; Kong, Y.L.; Malatesta, K.A.; Soboyejo, W.O.; Verma, N.; Gracias, D.H.; McAlpine, M.C. 3D
printed bionic ears. Nano Lett. 2013,13, 2634–2639. [CrossRef] [PubMed]
33.
Kang, H.-W.; Lee, S.J.; Ko, I.K.; Kengla, C.; Yoo, J.J.; Atala, A. A 3D bioprinting system to produce human-scale tissue constructs
with structural integrity. Nat. Biotechnol. 2016,34, 312–319. [CrossRef] [PubMed]
34.
Gu, Z.; Fu, J.; Lin, H.; He, Y. Development of 3D bioprinting: From printing methods to biomedical applications. Asian J. Pharm.
Sci. 2019,15, 529–557. [CrossRef] [PubMed]
35.
Hölzl, K.; Lin, S.; Tytgat, L.; Van Vlierberghe, S.; Gu, L.; Ovsianikov, A. Bioink properties before, during and after 3D bioprinting.
Biofabrication 2016,8, 032002. [CrossRef] [PubMed]
36.
Jiang, T.; Munguia-Lopez, J.G.; Flores-Torres, S.; Kort-Mascort, J.; Kinsella, J.M. Extrusion bioprinting of soft materials: An
emerging technique for biological model fabrication. Appl. Phys. Rev. 2019,6, 011310. [CrossRef]
37.
Blaeser, A.; Campos, D.D.; Puster, U.; Richtering, W.; Stevens, M.; Fischer, H. Controlling shear stress in 3D bioprinting is a key
factor to balance printing resolution and stem cell integrity. Adv. Healthc. Mater. 2016,5, 326–333. [CrossRef]
38. Irvine, S.A.; Venkatraman, S.S. Bioprinting and differentiation of stem cells. Molecules 2016,21, 1188. [CrossRef]
39. Stolberg, S.; McCloskey, K.E. Can shear stress direct stem cell fate? Biotechnol. Prog. 2009,25, 10–19. [CrossRef]
40.
Zhang, P.; Moudgill, N.; Hager, E.; Tarola, N.; DiMatteo, C.; McIlhenny, S.; Tulenko, T.; DiMuzio, P.J. Endothelial differentiation
of adipose-derived stem cells from elderly patients with cardiovascular disease. Stem Cells Dev.
2011
,20, 977–988. [CrossRef]
[PubMed]
41.
Colle, J.; Blondeel, P.; De Bruyne, A.; Bochar, S.; Tytgat, L.; Vercruysse, C.; Van Vlierberghe, S.; Dubruel, P.; Declercq, H. Bioprinting
predifferentiated adipose-derived mesenchymal stem cell spheroids with methacrylated gelatin ink for adipose tissue engineering.
J. Mater. Sci. Mater. Med. 2020,31, 36. [CrossRef]
42.
Leucht, A.; Volz, A.-C.; Rogal, J.; Borchers, K.; Kluger, P. Advanced gelatin-based vascularization bioinks for extrusion-based
bioprinting of vascularized bone equivalents. Sci. Rep. 2020,10, 5330. [CrossRef]
43.
Wang, X.-F.; Song, Y.; Liu, Y.-S.; Sun, Y.-c.; Wang, Y.-g.; Wang, Y.; Lyu, P.-J. Osteogenic differentiation of three-dimensional
bioprinted constructs consisting of human adipose-derived stem cells
in vitro
and
in vivo
.PLoS ONE
2016
,11, e0157214.
[CrossRef] [PubMed]
44.
Wehrle, M.; Koch, F.; Zimmermann, S.; Koltay, P.; Zengerle, R.; Stark, G.B.; Strassburg, S.; Finkenzeller, G. Examination of
hydrogels and mesenchymal stem cell sources for bioprinting of artificial osteogenic tissues. Cell. Mol. Bioeng.
2019
,12, 583–597.
[CrossRef]
45.
Yourek, G.; McCormick, S.M.; Mao, J.J.; Reilly, G.C. Shear stress induces osteogenic differentiation of human mesenchymal stem
cells. Regen. Med. 2010,5, 713–724. [CrossRef]
46.
Fedorovich, N.E.; De Wijn, J.R.; Verbout, A.J.; Alblas, J.; Dhert, W.J. Three-dimensional fiber deposition of cell-laden, viable,
patterned constructs for bone tissue printing. Tissue Eng. Part A 2008,14, 127–133. [CrossRef] [PubMed]
47.
Rottensteiner-Brandl, U.; Detsch, R.; Sarker, B.; Lingens, L.; Köhn, K.; Kneser, U.; Bosserhoff, A.K.; Horch, R.E.; Boccaccini, A.R.;
Arkudas, A. Encapsulation of rat bone marrow derived mesenchymal stem cells in alginate dialdehyde/gelatin microbeads with
and without nanoscaled bioactive glass for in vivo bone tissue engineering. Materials 2018,11, 1880. [CrossRef] [PubMed]
48.
Du, M.; Chen, B.; Meng, Q.; Liu, S.; Zheng, X.; Zhang, C.; Wang, H.; Li, H.; Wang, N.; Dai, J. 3D bioprinting of BMSC-laden
methacrylamide gelatin scaffolds with CBD-BMP2-collagen microfibers. Biofabrication 2015,7, 044104. [CrossRef] [PubMed]
49.
He, Y.; Gu, Z.; Xie, M.; Fu, J.; Lin, H. Why choose 3D bioprinting? Part II: Methods and bioprinters. Bio-Des. Manuf.
2020
,3, 1–4.
[CrossRef]
50.
Mandrycky, C.; Wang, Z.; Kim, K.; Kim, D.-H. 3D bioprinting for engineering complex tissues. Biotechnol. Adv.
2016
,34, 422–434.
[CrossRef]
51.
Kim, J.D.; Choi, J.S.; Kim, B.S.; Choi, Y.C.; Cho, Y.W. Piezoelectric inkjet printing of polymers: Stem cell patterning on polymer
substrates. Polymer 2010,51, 2147–2154. [CrossRef]
52.
Ru, J.; Guo, L.; Ji, Y.; Niu, Y. Hydrostatic pressure induces osteogenic differentiation of adipose-derived mesenchymal stem cells
through increasing lncRNA-PAGBC. Aging 2020,12, 13477. [CrossRef] [PubMed]
53.
Puetzer, J.; Williams, J.; Gillies, A.; Bernacki, S.; Loboa, E.G. The effects of cyclic hydrostatic pressure on chondrogenesis and
viability of human adipose-and bone marrow-derived mesenchymal stem cells in three-dimensional agarose constructs. Tissue
Eng. Part A 2013,19, 299–306. [CrossRef] [PubMed]
Sensors 2021,21, 7477 18 of 20
54.
Shav, D.; Einav, S. The effect of mechanical loads in the differentiation of precursor cells into mature cells. Ann. N. Y. Acad. Sci.
2010,1188, 25–31. [CrossRef] [PubMed]
55.
Gao, G.; Yonezawa, T.; Hubbell, K.; Dai, G.; Cui, X. Inkjet-bioprinted acrylated peptides and PEG hydrogel with human
mesenchymal stem cells promote robust bone and cartilage formation with minimal printhead clogging. Biotechnol. J.
2015
,10,
1568–1577. [CrossRef]
56.
Gao, G.; Schilling, A.F.; Hubbell, K.; Yonezawa, T.; Truong, D.; Hong, Y.; Dai, G.; Cui, X. Improved properties of bone and
cartilage tissue from 3D inkjet-bioprinted human mesenchymal stem cells by simultaneous deposition and photocrosslinking in
PEG-GelMA. Biotechnol. Lett. 2015,37, 2349–2355. [CrossRef] [PubMed]
57.
Dhawan, A.; Kennedy, P.M.; Rizk, E.B.; Ozbolat, I.T. Three-dimensional bioprinting for bone and cartilage restoration in
orthopaedic surgery. JAAOS-J. Am. Acad. Orthop. Surg. 2019,27, e215–e226. [CrossRef] [PubMed]
58.
wati Midha, S.; Patra, P.; Mohanty, S. Advances in 3D Bioprinting of Bone: Progress and Challenges. J. Tissue Eng. Regen. Med.
2019,13, 925–945. [CrossRef]
59.
Koch, L.; Kuhn, S.; Sorg, H.; Gruene, M.; Schlie, S.; Gaebel, R.; Polchow, B.; Reimers, K.; Stoelting, S.; Ma, N. Laser printing of skin
cells and human stem cells. Tissue Eng. Part C Methods 2010,16, 847–854. [CrossRef] [PubMed]
60.
Gruene, M.; Pflaum, M.; Hess, C.; Diamantouros, S.; Schlie, S.; Deiwick, A.; Koch, L.; Wilhelmi, M.; Jockenhoevel, S.; Haverich, A.
Laser printing of three-dimensional multicellular arrays for studies of cell–cell and cell–environment interactions. Tissue Eng.
Part C Methods 2011,17, 973–982. [CrossRef] [PubMed]
61.
Sorkio, A.; Koch, L.; Koivusalo, L.; Deiwick, A.; Miettinen, S.; Chichkov, B.; Skottman, H. Human stem cell based corneal tissue
mimicking structures using laser-assisted 3D bioprinting and functional bioinks. Biomaterials 2018,171, 57–71. [CrossRef]
62.
Somaiah, C.; Kumar, A.; Mawrie, D.; Sharma, A.; Patil, S.D.; Bhattacharyya, J.; Swaminathan, R.; Jaganathan, B.G. Collagen
promotes higher adhesion, survival and proliferation of mesenchymal stem cells. PLoS ONE 2015,10, e0145068.
63.
Gruene, M.; Pflaum, M.; Deiwick, A.; Koch, L.; Schlie, S.; Unger, C.; Wilhelmi, M.; Haverich, A.; Chichkov, B. Adipogenic
differentiation of laser-printed 3D tissue grafts consisting of human adipose-derived stem cells. Biofabrication
2011
,3, 015005.
[CrossRef] [PubMed]
64.
Clause, K.C.; Liu, L.J.; Tobita, K. Directed stem cell differentiation: The role of physical forces. Cell Commun. Adhes.
2010
,17,
48–54. [CrossRef] [PubMed]
65.
Gruene, M.; Deiwick, A.; Koch, L.; Schlie, S.; Unger, C.; Hofmann, N.; Bernemann, I.; Glasmacher, B.; Chichkov, B. Laser printing
of stem cells for biofabrication of scaffold-free autologous grafts. Tissue Eng. Part C Methods
2011
,17, 79–87. [CrossRef] [PubMed]
66.
Rémy, M.; Kériquel, V.; Correa, M.M.; Guillotin, B.; Guillemot, F. Creation of highly defined mesenchymal stem cell patterns in
three dimensions by laser-assisted bioprinting. J. Nanotechnol. Eng. Med. 2015,6, 021006.
67.
Ali, M.; Pages, E.; Ducom, A.; Fontaine, A.; Guillemot, F. Controlling laser-induced jet formation for bioprinting mesenchymal
stem cells with high viability and high resolution. Biofabrication 2014,6, 045001. [CrossRef]
68.
Keriquel, V.; Guillemot, F.; Arnault, I.; Guillotin, B.; Miraux, S.; Amédée, J.; Fricain, J.-C.; Catros, S.
In vivo
bioprinting for
computer-and robotic-assisted medical intervention: Preliminary study in mice. Biofabrication 2010,2, 014101. [CrossRef]
69.
Keriquel, V.; Oliveira, H.; Rémy, M.; Ziane, S.; Delmond, S.; Rousseau, B.; Rey, S.; Catros, S.; Amédée, J.; Guillemot, F. In situ
printing of mesenchymal stromal cells, by laser-assisted bioprinting, for
in vivo
bone regeneration applications. Sci. Rep.
2017
,7,
1778. [CrossRef]
70.
Billiet, T.; Gevaert, E.; De Schryver, T.; Cornelissen, M.; Dubruel, P. The 3D printing of gelatin methacrylamide cell-laden
tissue-engineered constructs with high cell viability. Biomaterials 2014,35, 49–62. [CrossRef]
71.
Campos, D.F.D.; Blaeser, A.; Weber, M.; Jäkel, J.; Neuss, S.; Jahnen-Dechent, W.; Fischer, H. Three-dimensional printing of stem
cell-laden hydrogels submerged in a hydrophobic high-density fluid. Biofabrication 2012,5, 015003. [CrossRef]
72.
Chang, C.C.; Boland, E.D.; Williams, S.K.; Hoying, J.B. Direct-write bioprinting three-dimensional biohybrid systems for future
regenerative therapies. J. Biomed. Mater. Res. Part B Appl. Biomater. 2011,98, 160–170. [CrossRef] [PubMed]
73.
Chang, R.; Nam, J.; Sun, W. Effects of dispensing pressure and nozzle diameter on cell survival from solid freeform fabrication–
based direct cell writing. Tissue Eng. Part A 2008,14, 41–48. [CrossRef]
74.
Gao, Q.; He, Y.; Fu, J.-z.; Liu, A.; Ma, L. Coaxial nozzle-assisted 3D bioprinting with built-in microchannels for nutrients delivery.
Biomaterials 2015,61, 203–215. [CrossRef] [PubMed]
75.
Hopp, B.; Smausz, T.; Kresz, N.; Barna, N.; Bor, Z.; Kolozsvári, L.; Chrisey, D.B.; Szabó, A.; Nógrádi, A. Survival and proliferative
ability of various living cell types after laser-induced forward transfer. Tissue Eng. 2005,11, 1817–1823. [CrossRef] [PubMed]
76.
Marga, F.; Jakab, K.; Khatiwala, C.; Shepherd, B.; Dorfman, S.; Hubbard, B.; Colbert, S.; Forgacs, G. Toward engineering functional
organ modules by additive manufacturing. Biofabrication 2012,4, 022001. [CrossRef] [PubMed]
77.
Melchels, F.P.; Dhert, W.J.; Hutmacher, D.W.; Malda, J. Development and characterisation of a new bioink for additive tissue
manufacturing. J. Mater. Chem. B 2014,2, 2282–2289. [CrossRef] [PubMed]
78.
Nair, K.; Gandhi, M.; Khalil, S.; Yan, K.C.; Marcolongo, M.; Barbee, K.; Sun, W. Characterization of cell viability during bioprinting
processes. Biotechnol. J. Healthc. Nutr. Technol. 2009,4, 1168–1177. [CrossRef] [PubMed]
79.
Smith, C.M.; Stone, A.L.; Parkhill, R.L.; Stewart, R.L.; Simpkins, M.W.; Kachurin, A.M.; Warren, W.L.; Williams, S.K. Three-
dimensional bioassembly tool for generating viable tissue-engineered constructs. Tissue Eng.
2004
,10, 1566–1576. [CrossRef]
[PubMed]
Sensors 2021,21, 7477 19 of 20
80.
Billiet, T.; Vandenhaute, M.; Schelfhout, J.; Van Vlierberghe, S.; Dubruel, P. A review of trends and limitations in hydrogel-rapid
prototyping for tissue engineering. Biomaterials 2012,33, 6020–6041. [CrossRef]
81.
Li, J.; Rossignol, F.; Macdonald, J. Inkjet printing for biosensor fabrication: Combining chemistry and technology for advanced
manufacturing. Lab Chip 2015,15, 2538–2558. [CrossRef] [PubMed]
82. Calvert, P. Inkjet printing for materials and devices. Chem. Mater. 2001,13, 3299–3305. [CrossRef]
83.
Cui, X.; Dean, D.; Ruggeri, Z.M.; Boland, T. Cell damage evaluation of thermal inkjet printed Chinese hamster ovary cells.
Biotechnol. Bioeng. 2010,106, 963–969. [CrossRef]
84.
Saunders, R.E.; Derby, B. Inkjet printing biomaterials for tissue engineering: Bioprinting. Int. Mater. Rev.
2014
,59, 430–448.
[CrossRef]
85.
Xu, T.; Jin, J.; Gregory, C.; Hickman, J.J.; Boland, T. Inkjet printing of viable mammalian cells. Biomaterials
2005
,26, 93–99.
[CrossRef]
86.
Ozbolat, I.T.; Yu, Y. Bioprinting toward organ fabrication: Challenges and future trends. IEEE Trans. Biomed. Eng.
2013
,60,
691–699. [CrossRef] [PubMed]
87.
Guillotin, B.; Guillemot, F. Cell patterning technologies for organotypic tissue fabrication. Trends Biotechnol.
2011
,29, 183–190.
[CrossRef]
88.
Guillotin, B.; Souquet, A.; Catros, S.; Duocastella, M.; Pippenger, B.; Bellance, S.; Bareille, R.; Rémy, M.; Bordenave, L.; Amédée, J.
Laser assisted bioprinting of engineered tissue with high cell density and microscale organization. Biomaterials
2010
,31, 7250–7256.
[CrossRef] [PubMed]
89.
Jia, J.; Richards, D.J.; Pollard, S.; Tan, Y.; Rodriguez, J.; Visconti, R.P.; Trusk, T.C.; Yost, M.J.; Yao, H.; Markwald, R.R. Engineering
alginate as bioink for bioprinting. Acta Biomater. 2014,10, 4323–4331. [CrossRef]
90.
Yang, Y.; Wang, X.; Wang, Y.; Hu, X.; Kawazoe, N.; Yang, Y.; Chen, G. Influence of cell spreading area on the osteogenic
commitment and phenotype maintenance of mesenchymal stem cells. Sci. Rep. 2019,9, 6891. [CrossRef]
91.
Kim, D.; Monaco, E.; Maki, A.; De Lima, A.S.; Kong, H.J.; Hurley, W.L.; Wheeler, M.B. Morphologic and transcriptomic comparison
of adipose-and bone-marrow-derived porcine stem cells cultured in alginate hydrogels. Cell Tissue Res.
2010
,341, 359–370.
[CrossRef]
92.
Engler, A.J.; Sen, S.; Sweeney, H.L.; Discher, D.E. Matrix elasticity directs stem cell lineage specification. Cell
2006
,126, 677–689.
[CrossRef] [PubMed]
93.
Guneta, V.; Loh, Q.L.; Choong, C. Cell-secreted extracellular matrix formation and differentiation of adipose-derived stem cells in
3D alginate scaffolds with tunable properties. J. Biomed. Mater. Res. Part A 2016,104, 1090–1101. [CrossRef] [PubMed]
94.
Ghiasi, M.; Kalhor, N.; Tabatabaei Qomi, R.; Sheykhhasan, M. The effects of synthetic and natural scaffolds on viability and
proliferation of adipose-derived stem cells. Front. Life Sci. 2016,9, 32–43. [CrossRef]
95.
Alsberg, E.; Anderson, K.; Albeiruti, A.; Franceschi, R.; Mooney, D. Cell-interactive alginate hydrogels for bone tissue engineering.
J. Dent. Res. 2001,80, 2025–2029. [CrossRef] [PubMed]
96. Lee, K.Y.; Mooney, D.J. Alginate: Properties and biomedical applications. Prog. Polym. Sci. 2012,37, 106–126. [CrossRef]
97.
Osathanon, T.; Giachelli, C.M.; Somerman, M.J. Immobilization of alkaline phosphatase on microporous nanofibrous fibrin
scaffolds for bone tissue engineering. Biomaterials 2009,30, 4513–4521. [CrossRef]
98.
Jafary, F.; Hanachi, P.; Gorjipour, K. Osteoblast differentiation on collagen scaffold with immobilized alkaline phosphatase. Int. J.
Organ Transplant. Med. 2017,8, 195.
99.
Kakudo, N.; Shimotsuma, A.; Miyake, S.; Kushida, S.; Kusumoto, K. Bone tissue engineering using human adipose-derived stem
cells and honeycomb collagen scaffold. J. Biomed. Mater. Res. Part A 2008,84, 191–197. [CrossRef] [PubMed]
100.
Chen, H.; Qian, Y.; Xia, Y.; Chen, G.; Dai, Y.; Li, N.; Zhang, F.; Gu, N. Enhanced osteogenesis of ADSCs by the synergistic effect of
aligned fibers containing collagen I. ACS Appl. Mater. Interfaces 2016,8, 29289–29297. [CrossRef]
101.
Kim, A.Y.; Kim, Y.; Lee, S.H.; Yoon, Y.; Kim, W.-H.; Kweon, O.-K. Effect of gelatin on osteogenic cell sheet formation using canine
adipose-derived mesenchymal stem cells. Cell Transplant. 2017,26, 115–123. [CrossRef] [PubMed]
102.
Yoon, Y.; Jung, T.; Afan Shahid, M.; Khan, I.U.; Kim, W.H.; Kweon, O.-K. Frozen-thawed gelatin-induced osteogenic cell sheets
of canine adipose-derived mesenchymal stromal cells improved fracture healing in canine model. J. Vet. Sci.
2019
,20, e63.
[CrossRef] [PubMed]
103.
Wofford, A.; Bow, A.; Newby, S.; Brooks, S.; Rodriguez, R.; Masi, T.; Stephenson, S.; Gotcher, J.; Anderson, D.E.; Campbell, J.
Human Fat-Derived Mesenchymal Stem Cells Xenogenically Implanted in a Rat Model Show Enhanced New Bone Formation in
Maxillary Alveolar Tooth Defects. Stem Cells Int. 2020,2020, 8142938. [CrossRef]
104.
Wang, C.-Y.; Hong, P.-D.; Wang, D.-H.; Cherng, J.-H.; Chang, S.-J.; Liu, C.-C.; Fang, T.-J.; Wang, Y.-W. Polymeric Gelatin Scaffolds
Affect Mesenchymal Stem Cell Differentiation and Its Diverse Applications in Tissue Engineering. Int. J. Mol. Sci.
2020
,21, 8632.
[CrossRef] [PubMed]
105.
Aguiari, P.; Leo, S.; Zavan, B.; Vindigni, V.; Rimessi, A.; Bianchi, K.; Franzin, C.; Cortivo, R.; Rossato, M.; Vettor, R. High glucose
induces adipogenic differentiation of muscle-derived stem cells. Proc. Natl. Acad. Sci. USA
2008
,105, 1226–1231. [CrossRef]
[PubMed]
106.
Chen, P.-Y.; Huang, L.L.; Hsieh, H.-J. Hyaluronan preserves the proliferation and differentiation potentials of long-term cultured
murine adipose-derived stromal cells. Biochem. Biophys. Res. Commun. 2007,360, 1–6. [CrossRef]
Sensors 2021,21, 7477 20 of 20
107.
Espandar, L.; Bunnell, B.; Wang, G.Y.; Gregory, P.; McBride, C.; Moshirfar, M. Adipose-derived stem cells on hyaluronic
acid–derived scaffold: A new horizon in bioengineered cornea. Arch. Ophthalmol. 2012,130, 202–208. [CrossRef] [PubMed]
108.
Yeo, M.; Lee, J.-S.; Chun, W.; Kim, G.H. An innovative collagen-based cell-printing method for obtaining human adipose stem
cell-laden structures consisting of core–sheath structures for tissue engineering. Biomacromolecules
2016
,17, 1365–1375. [CrossRef]
109.
Xu, W.; Hu, R.; Fan, E.; Han, D. Adipose-derived mesenchymal stem cells in collagen—hyaluronic acid gel composite scaffolds
for vocal fold regeneration. Ann. Otol. Rhinol. Laryngol. 2011,120, 123–130. [CrossRef]
110.
Amann, E.; Wolff, P.; Breel, E.; van Griensven, M.; Balmayor, E.R. Hyaluronic acid facilitates chondrogenesis and matrix deposition
of human adipose derived mesenchymal stem cells and human chondrocytes co-cultures. Acta Biomater.
2017
,52, 130–144.
[CrossRef]
111.
Fernandez, T.G.; Tenorio, A.; Campbell, K.T.; Silva, E.A.; Leach, K. Evaluation of Alginate-Based Bioinks for 3D Bioprinting,
Mesenchymal Stromal Cell Osteogenesis, and Application for Patient-Specific Bone Grafts. bioRxiv 2020. [CrossRef]
112.
Scott, C.D.; Woodward, C.A.; Thompson, J.E. Solute diffusion in biocatalyst gel beads containing biocatalysis and other additives.
Enzym. Microb. Technol. 1989,11, 258–263. [CrossRef]
113.
Chan, W.W.; Yeo, D.C.L.; Tan, V.; Singh, S.; Choudhury, D.; Naing, M.W. Additive biomanufacturing with collagen inks.
Bioengineering 2020,7, 66. [CrossRef] [PubMed]
114.
Mizuno, M.; Kuboki, Y. Osteoblast-related gene expression of bone marrow cells during the osteoblastic differentiation induced
by type I collagen. J. Biochem. 2001,129, 133–138. [CrossRef]
115.
George, J.; Kuboki, Y.; Miyata, T. Differentiation of mesenchymal stem cells into osteoblasts on honeycomb collagen scaffolds.
Biotechnol. Bioeng. 2006,95, 404–411. [CrossRef]
116.
Chiu, L.-H.; Lai, W.-F.T.; Chang, S.-F.; Wong, C.-C.; Fan, C.-Y.; Fang, C.-L.; Tsai, Y.-H. The effect of type II collagen on MSC
osteogenic differentiation and bone defect repair. Biomaterials 2014,35, 2680–2691. [CrossRef] [PubMed]
117.
Mazaki, T.; Shiozaki, Y.; Yamane, K.; Yoshida, A.; Nakamura, M.; Yoshida, Y.; Zhou, D.; Kitajima, T.; Tanaka, M.; Ito, Y. A
novel, visible light-induced, rapidly cross-linkable gelatin scaffold for osteochondral tissue engineering. Sci. Rep.
2014
,4, 4457.
[CrossRef] [PubMed]
118.
Yao, M.; Zhang, J.; Gao, F.; Chen, Y.; Ma, S.; Zhang, K.; Liu, H.; Guan, F. New BMSC-laden gelatin hydrogel formed in situ by
dual-enzymatic cross-linking accelerates dermal wound healing. ACS Omega 2019,4, 8334–8340. [CrossRef] [PubMed]
119.
Banfi, G.; Iorio, E.L.; Corsi, M.M. Oxidative stress, free radicals and bone remodeling. Clin. Chem. Lab. Med.
2008
,46, 1550–1555.
[CrossRef] [PubMed]
120.
Zhu, M.; Lin, S.; Sun, Y.; Feng, Q.; Li, G.; Bian, L. Hydrogels functionalized with N-cadherin mimetic peptide enhance osteogenesis
of hMSCs by emulating the osteogenic niche. Biomaterials 2016,77, 44–52. [CrossRef]
121.
Cavallo, C.; Desando, G.; Ferrari, A.; Zini, N.; Mariani, E.; Grigolo, B. Hyaluronan scaffold supports osteogenic differentiation of
bone marrow concentrate cells. J. Biol. Regul. Homeost. Agents 2016,30, 409–420. [PubMed]
122.
Perez, R.A.; Kim, M.; Kim, T.-H.; Kim, J.-H.; Lee, J.H.; Park, J.-H.; Knowles, J.C.; Kim, H.-W. Utilizing core–shell fibrous
collagen-alginate hydrogel cell delivery system for bone tissue engineering. Tissue Eng. Part A
2014
,20, 103–114. [CrossRef]
[PubMed]
123.
Matsiko, A.; Levingstone, T.J.; O’Brien, F.J.; Gleeson, J.P. Addition of hyaluronic acid improves cellular infiltration and promotes
early-stage chondrogenesis in a collagen-based scaffold for cartilage tissue engineering. J. Mech. Behav. Biomed. Mater.
2012
,11,
41–52. [CrossRef] [PubMed]
124.
Zhang, Y.; Cao, Y.; Zhao, H.; Zhang, L.; Ni, T.; Liu, Y.; An, Z.; Liu, M.; Pei, R. An injectable BMSC-laden enzyme-catalyzed
crosslinking collagen-hyaluronic acid hydrogel for cartilage repair and regeneration. J. Mater. Chem. B
2020
,8, 4237–4244.
[CrossRef] [PubMed]
125.
Li, H.; Qi, Z.; Zheng, S.; Chang, Y.; Kong, W.; Fu, C.; Yu, Z.; Yang, X.; Pan, S. The application of hyaluronic acid-based hydrogels
in bone and cartilage tissue engineering. Adv. Mater. Sci. Eng. 2019,2019, 3027303. [CrossRef]
126.
Ilyas, A.; Lavrik, N.V.; Kim, H.K.; Aswath, P.B.; Varanasi, V.G. Enhanced interfacial adhesion and osteogenesis for rapid
“bone-like” biomineralization by PECVD-based silicon oxynitride overlays. ACS Appl. Mater. Interfaces
2015
,7, 15368–15379.
[CrossRef]
127.
Ilyas, A.; Odatsu, T.; Shah, A.; Monte, F.; Kim, H.K.; Kramer, P.; Aswath, P.B.; Varanasi, V.G. Amorphous silica: A new antioxidant
role for rapid critical-sized bone defect healing. Adv. Healthc. Mater. 2016,5, 2199–2213. [CrossRef]
