RESEARCH ARTICLE
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A Perfusable Multi-Hydrogel Vasculature On-ChipEngineered by 2-Photon 3D Printing and Scaﬀold Molding
to Improve Microfabrication Fidelity in Hydrogels
Federico Cantoni, Laurent Barbe, Hannah Pohlit, and Maria Tenje*
Compared to conventional cell culture, microphysiological systems (MPS),
also known as organs-on-chips, provide more precise control over thecell microenvironment, exhibit morebiomimetic cell culture conditions[2,3]and pave the way to a better understanding of biological mechanisms
including barrier function,[4] tumor
development[5] and drug transport.[6–8],
A key to developing in vitro 3D cell
models is the capacity to replicate theextracellular matrix (ECM) of the native
tissue with respect to mechanical andchemical properties.[9,10] Hydrogels,
which are polymer-based structures containing a large fraction of water, oﬀer avaluable tool to recreate human physiology in ways not possible with traditional
cell culture substrates.[11] The recent
development of synthetic and hybrid
polymers has overcome the batch-tobatch variation of natural hydrogels withmaintained cell responsive functionalities combined with mechanical stability
and good processability.[12–14] However,these hydrogels still exhibit volume changes when equilibratingwith a cell culture medium, which can then hamper microstructure printing ﬁdelity.[15,16]
One of the current challenges in 3D cell culture is to build vascular systems into the 3D culture scaﬀolds to ensure nutrient exchange and waste removal, and therefore prevent cell necrosis in
the bulk of the material.[17]A widely used approach to generate perfusable vascular networks relies on the biological process of angiogenesis within the
microﬂuidic device.[18–21] The process mimics in vivo architecture and dimensions, but the poor control on the vessel density and geometry hinders comparative studies. To achieve better
control, an engineering approach to fabricate perfusable vascular
networks is taken. This can include the casting of hydrogel precursor solution around sacriﬁcial structures and needles,[22,23] orforming vessels via the so-called viscous ﬁngering method.[24,25]
Although the simplicity of these techniques is attractive, the freedom in design and spatial positioning as well as the required resolution for the reproduction of in vivo tissue architecture features
is limited.

Engineering vasculature networks in physiologically relevant hydrogels
represents a challenge in terms of both fabrication, due to the cell–bioink
interactions, as well as the subsequent hydrogel-device interfacing. Here, a
new cell-friendly fabrication strategy is presented to realize perfusable
multi-hydrogel vasculature models supporting co-culture integrated in a
microﬂuidic chip. The system comprises two diﬀerent hydrogels to speciﬁcallysupport the growth and proliferation of two diﬀerent cell types selected for thevessel model. First, the channels are printed in a gelatin-based ink by
two-photon polymerization (2PP) inside the microﬂuidic device. Then, ahuman lung ﬁbroblast-laden ﬁbrin hydrogel is injected to surround the printed
network. Finally, human endothelial cells are seeded inside the printed
channels. The printing parameters and ﬁbrin composition are optimized toreduce hydrogel swelling and ensure a stable model that can be perfused withcell media. Fabricating the hydrogel structure in two steps ensures that nocells are exposed to cytotoxic fabrication processes, while still obtaining highﬁdelity printing. In this work, the possibility to guide the endothelial cell
invasion through the 3D printed scaﬀold and perfusion of the co-culture
model for 10 days is successfully demonstrated on a custom-made perfusion
system.

1. Introduction
In vitro cell-based models are important tools in drug development and medical research. Traditional 2D cell culture using
Petri dishes or Transwell inserts does however suﬀer from a nonphysiological environment that may alter the cell behavior and in
turn, aﬀect the experimental outcomes.[1]F. Cantoni, L. Barbe, H. Pohlit, M. Tenje
Department of Materials Science and Engineering
Science for Life Laboratory
Uppsala University
Box 35, Uppsala 751 03, Sweden
E-mail: maria.tenje@angstrom.uu.se
The ORCID identiﬁcation number(s) for the author(s) of this article
can be found under https://doi.org/10.1002/admt.202300718© 2024 The Authors. Advanced Materials Technologies published by
Wiley-VCH GmbH. This is an open access article under the terms of theCreative Commons Attribution License, which permits use, distribution
and reproduction in any medium, provided the original work is properly
cited.

DOI: 10.1002/admt.202300718

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Physiologically relevant 3D vasculature networks have beenobtained by combining casting with more sophisticated 3D
printed sacriﬁcial templates.[26,27] Alternatively, multi-material
extrusion through co-axial needles can be used to create hollow
structures[28] with the possibility to generate co-cultures.[29] However, such techniques are unable to provide a resolution <100 μm,
and exhibit poor compatibility with on-chip fabrication as theyrequire rather complex manipulations to integrate the hydrogel
scaﬀold.[30]A replica vasculature network model can be created on-chip by
UV-assisted stereolithography 3D printing that provides a resolution of ≈ 5 μm and cell-friendly crosslinking chemistry.[31–35]
Despite the technology being suitable for on-chip fabrication,
the reduced ability to print hollow structures on-chip with anaxial resolution <100 μm prevents the technology from being
used for MPS.[36] Whilst the recently developed volumetric 3D
printing presents unprecedented printing speed and surface
quality,[37–39] it still displays a resolution and minimal featuresize more suitable for hollow structures >400 μm.[40,41] 4D printing shows promise in addressing the resolution limits by creating a self-folding tubular structure caused by inhomogeneous
swelling of a planar hydrogel scaﬀold.[42,43] However, the technique poses challenges in creating ramiﬁed hollow structuresand further developments are required to achieve sub-100 μm
resolution.[44]Two-photon polymerization technology has recently emerged
as a high-ﬁdelity 3D printing technique capable of generating
hollow structures by conﬁning crosslinking to the laser voxel
via a non-linear absorption process.[45–50] The achievable submicron feature size and resolution are suitable for cell-scale studies
including scaﬀold porosity,[51,52] mechanical gradient eﬀects[53],and cell-substrate surface interaction.[54,55] Despite the recent development of novel water-soluble and biocompatible chemistry,
the technology however still faces a limited portfolio of biocompatible materials, and the photo-initiator radical generation isa hazard for cells.[56,57] In addition, the lengthy fabrication requires the sample to be placed in an incubator-like environment
during the full printing process to preserve cell viability[58–60]
with the drawback of cell sedimentation.[61] Finally, the presence of cells in the bioink negatively aﬀects polymer chain
crosslinking due to light scattering and undesired interactions
with the precursor components, thus compromising the printing
ﬁdelity.[62–64]
To circumvent the photo-initiator toxicity, 2-photon ablationand cavitation have been investigated for generating channel networks in collagen hydrogels.[65–68] Such techniques provide an alternative process that still matches the resolution of 2PP with theadvantage of being less harmful to the pre-encapsulated cells dueto the smaller processing area.[69] However, the cell concentrationand hydrogel turbidity interfere with the printing quality and theissue with a long process time still remains.[70]In our work, we propose a new strategy to form high-ﬁdelity
printing of complex vascular channels inside natural cell-ladenpolymers without causing cell damage during fabrication. Thisapproach builds upon a two-step process that ﬁrst sees the fabrication of a hydrogel channel network by 2PP directly inside a
microﬂuidic device. After generating the vasculature backbone,
another cell-laden hydrogel can be injected to surround theprinted structures. Finally, endothelial cells can be seeded inside

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the printed hydrogel channels to form a vascularized system.
The proposed technique is capable of creating a multi-hydrogel
construct to accommodate diﬀerent cell needs in the case ofa co-culture system. All steps are performed directly insidea PDMS/glass microﬂuidic device, ensuring a user-friendly
process and the possibility to directly connect the vascular
network with external pumps for cell media perfusion. Here,
2PP-structured gelatin-based hydrogels with feature sizes down
to 10 μm displayed a stable scaﬀold and enabled co-culture ofhuman umbilical vein endothelial cell (HUVEC)/ﬁbroblastladen ﬁbrin over 10 days in a customized perfusion
system.

2. Results and Discussion
An ideal bio-ink should provide appropriate biocompatibility forthe cultured cells to ensure the formation of functional tissues.
Encapsulating the cells in the hydrogel precursor solution makes
this process more challenging as it requires a biocompatible fabrication process. It also has a negative impact on the crosslinking mechanisms, which reduces printing quality.[63] At the sametime, hydrogels used as 3D cell culture scaﬀolds must also exhibit
suﬃcient mechanical stability to support cell media perfusion
and withstand the fabrication process.[11,70] These requirements
combined make the synthesis of suitable hydrogel polymers andthe development of fabrication techniques challenging.[61,71]
To overcome these problems, we combined high-resolution
2PP printing of a biocompatible ink with a second hydrogel surrounding the structures inside a microﬂuidic chip. This takes advantage of combining two diﬀerent hydrogels, each compatible
with one cell type (Figure 1). Since previous studies have demonstrated that gelatin-derived hydrogels are a suitable substrate for
HUVEC culture, a 2PP gelatin-based hydrogel was chosen forthe high-resolution printing of the vasculature backbone.[49,72,73]
The chosen 2PP hydrogel displays mechanical properties between 0.5–2 kPa (data provided by the supplier), comparable withthe stiﬀness range for HUVEC culture as shown in previous
studies.[66,68,74] Fibrin was used for the encapsulation of ﬁbroblasts as it has previously been demonstrated to support the proliferation of human lung ﬁbroblasts (HLFs).[75,76]
The presented fabrication strategy and chip design have beendeveloped to address four main challenges in high-resolution
3D bioprinting: cytotoxicity, media perfusion, lengthy fabrication, and printing ﬁdelity. When conventional high-resolution 3D
printing is used for cell-laden hydrogels, two main negative effects occur: radical formation during light-based printing, andhigh shear stress in the case of extrusion systems.[56,69] This 2step strategy prevents cell damage from these sources as cells are
not included during the actual printing process.
To address the challenge of media perfusion, the hydrogel
structure was fabricated inside a microﬂuidic platform. This offers a reliable hydrogel-device interface for cell seeding and cell
media perfusion while still ensuring high-resolution imaging.[77]Printing of the vasculature backbone without encapsulated
cells allowed high printing ﬁdelity as cell scattering could not affect the fabrication process.[63] In addition, printing the hydrogel
channels with wall thicknesses down to 40 μm reduced the inﬂuence of gelatin-derived hydrogel swelling on the printing quality,
while also reducing the printing time of the whole scaﬀold.[78]© 2024 The Authors. Advanced Materials Technologies published by Wiley-VCH GmbH2365709x, 2024, 4, Downloaded from https://onlinelibrary.wiley.com/doi/10.1002/admt.202300718 by Uppsala University Library, Wiley Online Library on [22/05/2024]. See the Terms and Conditions (https://onlinelibrary.wiley.com/terms-and-conditions) on Wiley Online Library for rules of use; OA articles are governed by the applicable Creative Commons License

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Figure 1. Schematic representation of the cell-friendly fabrication strategy for perfusable co-culture of HUVECs and ﬁbroblasts. a) The empty microﬂuidic
chip and the magniﬁcation in the coverslip holes. b) Injection of the 2PP ink and printing of the hydrogel channel. c) Injection of ﬁbroblast-laden ﬁbrinsurrounding the hydrogel vasculature and d) seeding of HUVECs inside the 2PP hydrogel channel connected for cell media perfusion.

2.1. Printing Optimization and Channel Formation
2.1.1. Printing Parameter Screening to Identify the Minimal
Hydrogel Swelling
The most critical aspect when printing high-resolution structures
is to ensure the highest printing ﬁdelity. Despite hydrogels being 3D crosslinked polymer structures, the matrix is still susceptible to volume change over at least 24 h following printing[35]as the matrix equilibrates in aqueous solutions. We therefore investigated the eﬀect of printing parameters on hydrogel swellingin cell media with the aim of minimizing swelling. A parameter screening, including printing speed (200–1000 mm s−1 at
200 mm s−1 intervals) and slicing layer (1–9 μm at 2 μm intervals) was performed with 0.5 μm constant hatching (Figure 2).
The slicing parameters “top-down” and “woodpile mode” werechosen over “one-direction printing” and “bottom-up mode”, respectively, since these have been reported to provide better printing ﬁdelity in previous studies.[49] After printing, all structures
were incubated in cell media for 24 h to reach equilibrium.
No structures were retained on the surface for the 9 μm slicing layer, and a printing speed of 1000 mm s−1 resulted in heavily distorted structures due to the non-synchronization between
the laser on/oﬀ switching and the galvanic scanning mirror(Figure 2). The pillars with a 7 μm slicing layer exhibited partiallyprinted structures and were not always retained on the glass surface, while the 1 μm slicing layer resulted in distorted structures

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that displayed rounded corners and a signiﬁcantly deformed
shape when compared to the other slicing layers (Video S1, Supporting Information). Since the stability of the DTT crosslinker
reduces as the temperature increases,[79] we speculate that the
1 μm slicing layer results in too much energy being supplied perunit volume, causing an excessive localized increase in temperature. This hypothesis is supported by observing the increase in
deformation and swelling as the printing speed decreases, which
resulted in an even higher power localization, and in turn, increased temperature. The inactivation of DTT causes a depletion of the crosslinker, which in turn aﬀects the crosslinker-tofunctional group ratio and therefore the hydrogel crosslinking.
Then, as reported in previous studies, the high energy density can
trigger a fast localized cross-linking, drastically increasing the viscosity and reducing the diﬀusion of unreacted DTT molecules to
the gelatin sites, which also results in a decrease in crosslinking
density.[53,80]
Consequently, only samples sliced with 3 and 5 μm intervals
and scanning speeds ranging between 200 and 800 mm s−1 were
further analyzed to measure the top surface area (Figure S1, Supporting Information) and calculate the linear swelling (Figure 2b)with statistical analysis (Table S1, Supporting Information).
The results for the 5 μm slicing indicated a constant increasein swelling toward higher printing speeds as the energy dose decreased with the increase in scanning speed. The swelling observed in the 3 μm slicing was generally smaller than the 5 μm
slicing, but diﬀered in relation to the printing speed. For the© 2024 The Authors. Advanced Materials Technologies published by Wiley-VCH GmbH2365709x, 2024, 4, Downloaded from https://onlinelibrary.wiley.com/doi/10.1002/admt.202300718 by Uppsala University Library, Wiley Online Library on [22/05/2024]. See the Terms and Conditions (https://onlinelibrary.wiley.com/terms-and-conditions) on Wiley Online Library for rules of use; OA articles are governed by the applicable Creative Commons License

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Figure 2. a) Fluorescent image showing resulting hydrogel dimensions when varying slicing interlayer distance and scanning speed of the 2PP processusing constant power of 100 mW and constant hatching of 0.5 μm. b) Linear swelling for structures prepared using either 3 or 5 μm layer spacing. Errorbar from standard deviation (n = 3).

3 μm slicing samples, a decrease in swelling for printing speed
between 200 and 600 mm s−1 and an increase in swelling from
600 mm s−1 was observed. From this study, we concluded thatprinter settings of 600 mm s−1 and 3 μm slicing were optimal
to minimize hydrogel swelling and these settings were usedthroughout the manuscript.
To ensure the observed size diﬀerence between the intended
design and obtained structure was caused by hydrogel swellingand not poor printing capability of the printer itself, we measured
the dimension of a hydrogel layer in contact with glass, that is,
at a position where it is unable to change dimensions due to surface constraints. An average deviation from the CAD value of only
4% was measured, indicating that hydrogel swelling is the maincause of size deviations in the printed structures reported above.

2.1.2. The Double Challenge in Bioprinting: High Cell Viability with
Good Printing Fidelity
A system supporting cells co-cultured in a hydrogel scaﬀold withan integrated vasculature needs to support both the culture ofendothelial cells on the channel lumen’s surface and the cells inthe scaﬀold bulk. Consequently, we investigated the ability of the
2PP ink to support 2D culture of HUVECs on the surface of theprinted scaﬀold (Figure 3a–d), and 3D culture of HLFs encapsulated in the hydrogel matrix (Figure 3d–f).
The biocompatibility of the 2PP hydrogel for 2D culture wasinvestigated by culturing HUVECs on the surface of 2PP-printedhydrogel disks. HUVECs cultured on a glass surface coated withan adhesion-promoting solution (Speed Coating, PELOBiotech)
was used as control. For both conditions, high cell viability wasobserved after 10 days. The control experiment displayed a higher
viability of ≈95% against the 85% shown for the hydrogel scaﬀold(Figure 3d).
In addition to biocompatibility, proliferation was also investigated to evaluate the ability of cells to populate the scaﬀold.

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Steady growth of the HUVECs was seen for the ﬁrst 7 days
(Figure 3c). The cells on the hydrogel substrate showed a slower
proliferation on day 1 but this did not result in a lower proliferation over the full 10-day-period. No hydrogel detachment or hydrogel bending was observed on day 10 and a z-stack video of thecells on the 2PP hydrogel inside the device is shown (Video S2,
Supporting Information). The microﬂuidic chip enabled hydrogel disk printing as well as cell culture, staining, and cell imaging.
The miniaturization of the wells on-chip oﬀered a valid screening platform for 2PP inks as conventional well plates present too
big surfaces for 2-photon 3D printing (Figure 3b).
The possibility of encapsulating cells in the 2PP ink was evaluated with HLFs. A ﬁbroblasts-laden 2PP ink was printed in cube
samples (500 × 500 × 250 μm3 ) using the optimized printing parameters identiﬁed in Section 2.1.1. Fibroblast-laden ﬁbrin hydrogels served as a control for the biocompatibility assay since ﬁbrin is commonly used for ﬁbroblast culture.[81,82] No signiﬁcantdiﬀerence in ﬁbroblast viability was observed between the two different hydrogels, with both showing values ≈ 90%, (Figure 3d).
However, a cell eﬀect was observed on the printing quality of
the cell-laden structure, with the 2PP ink exhibiting areas withlower hydrogel signal surrounding the cells (Figure 3e,f; Video
S3, Supporting Information). The lower hydrogel signal was associated with reduced scaﬀold crosslinking due to the cells acting as a source of light scattering and termination of the free
radicals.[83,84] This has previously been reported as a major issue
with 2PP of cell-laden hydrogels, in which the cells aﬀect printing ﬁdelity by creating non-uniform crosslinking and areas in the
structure with weak mechanical properties.[62,64]
As a possible solution, crosslinking of the aﬀected area would
beneﬁt from a higher energy dosage. However, higher energies
would negatively inﬂuence the cell viability[53,59] and also drift
away from the optimized printing parameters, shown in Section 2.1.1, and cause a higher degree of swelling. In addition, excessive energy dosages are not suitable in the case of sub 100 μm
hollow structures, since over-polymerization can occur, leading© 2024 The Authors. Advanced Materials Technologies published by Wiley-VCH GmbH2365709x, 2024, 4, Downloaded from https://onlinelibrary.wiley.com/doi/10.1002/admt.202300718 by Uppsala University Library, Wiley Online Library on [22/05/2024]. See the Terms and Conditions (https://onlinelibrary.wiley.com/terms-and-conditions) on Wiley Online Library for rules of use; OA articles are governed by the applicable Creative Commons License

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Figure 3. Biocompatibility of 2PP hydrogel for 2D and 3D cultures. a) Schematic representation of the 2PP hydrogel scaﬀold fabrication for seeding of
HUVECs. b) Optical micrograph of the device with cells growing on the hydrogel, blue: 2PP ink, green: GFP-HUVECs, red: Cell nuclei. c) Comparison
between the cell number over time when cultured on the 2PP hydrogel scaﬀold and a glass surface coated with speed coating. Error bars from standard
deviation (n = 9). d) Viability for 2D and 3D culture of HUVECs and ﬁbroblasts with the 2PP hydrogel after 24 h, respectively (n = 5). e) Optical micrographshowing the top view and side cross-sections of a ﬁbroblast-laden hydrogel, blue: 2PP ink, green: Fibroblasts. f) Signal intensity along the horizontal
cross-section (red rectangle in image d)) for the hydrogel (blue) and the cell (green).

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www.advmattechnol.deto occlusions of channels and chambers.[53] By combining 2PP
and micromolding, we address the printing limitations causedby the presence of cells.
Despite fabrication being performed outside an incubator, theencapsulated ﬁbroblast displayed a high viability. In addition,
sedimentation of cells at the bottom of the microﬂuidic platform
was observed during 3D printing, causing a non-uniform cell distribution in the printed structure. The conventional strategies of
reducing sedimentation by either adding a viscosity enhancer[85]or applying gentle movement to the resin[86] are incompatible
due to their eﬀect on light absorption and printing ﬁdelity, respectively. From this study, we conclude that the 2PP bioink successfully supports 2D culture of HUVECs and that cell encapsulation entails limitations for high-printing ﬁdelity. Hence, we
utilized ﬁbrin for the 3D culture gel surrounding the pre-deﬁned
vasculature scaﬀold obtained by 2PP.

2.1.3. The Fabrication Process Supports a Range of Hydrogel
Combinations
In photo-crosslinkable bioinks, the interaction between the cells
and the hydrogel precursor solution often has a negative impact
on biocompatibility and structure ﬁdelity.[62] These aspects combined with the hydrogel’s volume change upon reaching equilibrium with the cell medium still represent outstanding challenges
in high-resolution 3D bioprinting.
We propose a 2-step fabrication process to obtain multihydrogel structures on-chip that is cell-friendly with high 3D
printing ﬁdelity. This strategy avoids the inﬂuence of cells on
the fabrication process. First, a straight hydrogel channel was
printed with a 40 μm-thick wall, with a printing time of only
10 min. The 2PP printed design includes two hydrogel connectors with corresponding holes in the glass (Figure 4a,b). The hydrogel connectors present a wall thickness ≈175 μm to guarantee good adhesion with the glass and compensate for any defects in the hole, such as glass chipping generated during thelaser cutting. The 2PP scaﬀold height and width were smaller
than the dimension of the microﬂuidic channel to enable hydrogel swelling without constraint and to reduce deformations(Figure S2, Supporting Information). Then, a second hydrogel,here ﬁbrin, was manually injected by pipetting the precursor solution into the microﬂuidic device to fully surround the printed
structures. Injecting a second hydrogel to surround the vasculature network provided both a basis for 3D cell culture of a second cell type as well as a mechanical support for the printed hydrogel channel. Here, ﬁbrinogen and thrombin mixtures (2.5, 5,
and 10 mg mL−1 ﬁbrinogen with 0.5 U mL−1 thrombin) were
investigated to identify a suitable ﬁbrin precursor composition.
The 2.5 mg mL−1 ﬁbrinogen hydrogel retracted in areas close
to the connectors and along the channel structure after the removal of the non-crosslinked ink inside the channel and initiation of cell media perfusion, (Figure 4c–e), whereas the 5 and
10 mg mL−1 ﬁbrinogen hydrogels demonstrated good adhesion
to the whole channel structure. Both the 5 and 10 mg mL−1 ﬁbrinogen hydrogels resulted in perfusable channels, as conﬁrmed
by injecting 4 μm beads in the 2PP hydrogel channels and imaging with a confocal microscope in ﬂuorescence mode (Videos
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lar result, the 5 mg mL−1 ﬁbrinogen hydrogels were preferredover the 10 mg mL−1 ﬁbrinogen since better migration and proliferation of HLFs in this gel concentration has previously beenreported.[81]The second hydrogel can be chosen from a range of suitable hydrogels for 3D cell culture, depending on the application requirements. The versatility of the fabrication strategy was successfully
investigated by surrounding the printed vasculature with two
more hydrogels: pure collagen (2 mg mL−1 ) and a collagen/ﬁbrin
mixture (0.5 mg mL−1 collagen with 4.5 mg mL−1 ﬁbrin) precursor solution (Figure S3, Supporting Information).
Unlike light-based bioprinting, the proposed two-step strategy
enabled the fabrication of a vasculature-like system embedded in
diﬀerent natural hydrogels (ﬁbrin with diﬀerent ﬁbrinogen concentrations, collagen, and ﬁbrin–collagen) without the need tochange the process parameters and without any cell scattering
eﬀect.
The demonstrated approach of adding a second hydrogel tosurround the 2PP fabricated vessel also had additional positiveeﬀects. First, it provides mechanical support so that it is suﬃcient to make the wall of the vessel only 40 μm thick, which avoids
structure deformation due to the limited swelling. The thin vessel
wall reduces the printing time as the printing volume is conﬁned
to the connector and the 40 μm vessel wall. Structures obtainedwith a 200 μm-thick wall without ﬁbrin support displayed clogging of the channel due to signiﬁcant swelling of the 2PP hydrogel, (Figure S4, Supporting Information).
Most of the solutes and gas exchanges of the humanbody happen in the microvasculature,[87] inﬂuencing a large
number of physiological and pathological processes including angiogenesis,[88] barrier function,[89] inﬂammation, andthrombosis.[90] Thus, a fabrication strategy that enables the realization of in vitro models with feature sizes from only a few
micrometers up to 100 μm could contribute to understanding themechanisms in key biological events.
Using optimal 2PP conditions, inner diameters ranging from10 to 60 μm with a constant 40 μm wall thickness were printed toevaluate the ability of the technique to replicate tissue-like featuresizes.
As one of the main challenges of micro-scale hollow hydrogel
structures is to preserve the stability of the scaﬀold,[77,91] the inner diameter and eccentricity of the channels were measured at
24 h and 10 days after printing. The structure with the screened
diameters is shown in (Figure 5a–c).
All diameters were successfully printed and no collapse wasobserved even after 10 days of incubation. However, a change
in the inner geometry was noticed over the course of the experiment for all printed channels (Figure 5d). 24 h after printing,the smaller inner diameters (<40 μm) were closer in terms ofsize and eccentricity to the CAD design. Speciﬁcally, the channels with inner diameters ≥40 μm resulted in compression along
the y-axis, with an eccentricity between 0.7 and 0.9 (Figure 5d,e).
The change in eccentricity was most likely due to pressure applied during the injection of the ﬁbrin hydrogel. The vessels withlarger inner diameters proved to be mechanically weaker, resulting in structures more prone to deformation upon injection of thesecond hydrogel. Between days 1 and 10, an increase in the innerdiameter was observed in all evaluated channel structures. We hypothesized that the diameter change was due to the ﬁbrin matrix© 2024 The Authors. Advanced Materials Technologies published by Wiley-VCH GmbH2365709x, 2024, 4, Downloaded from https://onlinelibrary.wiley.com/doi/10.1002/admt.202300718 by Uppsala University Library, Wiley Online Library on [22/05/2024]. See the Terms and Conditions (https://onlinelibrary.wiley.com/terms-and-conditions) on Wiley Online Library for rules of use; OA articles are governed by the applicable Creative Commons License

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Figure 4. Optical images of the perfusable 3D channel were obtained by surrounding the 2PP structure with ﬁbrin precursor solutions with constantthrombin concentration (0.5 U mL−1 ) and diﬀerent ﬁbrinogen concentrations (2.5, 5, 10 mg mL−1 ). a) PDMS/glass microﬂuidic device used for structure
generation and perfusion. b) Microﬂuidic chip after 2PP printing (blue channel) and injection of the ﬁbrin. c) Sample obtained with 2.5 mg mL−1
ﬁbrinogen and 0.5 U mL−1 thrombin. The arrows indicate retraction areas of the ﬁbrin hydrogel after non-crosslinked ink removal. d) Sample obtainedwith 5 mg mL−1 ﬁbrinogen and 0.5 U mL−1 thrombin. e) Sample obtained with 10 mg mL−1 ﬁbrinogen and 0.5 U mL−1 thrombin. Red: ﬁbrin by reﬂection
mode. Blue: 2PP ink. Sample e) shows contaminants located mainly on the top part of the channel that are visible in reﬂection mode.

rearrangement caused by inelastic deformation during perfusion
and washing of the dye over the 10 days.[92] However, no physical damage was detected in the 2PP structures themselves as noleakage of Antonina red was noticed in the channel.
The technique allowed channels with inner diameters down to10 μm to be fabricated. Diameters smaller than this resulted in
clogged channels. In addition, walls thinner than 40 μm caused
the collapse of the structure when injecting the second hydrogel.
For further biological studies, channels with an inner diameter>30 μm were preferred to avoid too high shear stress for the cultured cells during perfusion and to simplify cell seeding.[93]2.1.4. The 2PP Hydrogel is Permeable to Molecules up to 250 kDa
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in several biological functions; regulating viability, migration,and proliferation of the cells, as well as nutrient exchange andwaste removal.[14] Speciﬁcally, ﬁbroblasts support HUVECs’ lumen formation by producing cell signaling molecules and ECM
components[94] (Table S2, Supporting information). Therefore, it
is important that the 2PP-deﬁned hydrogel channel is suﬃciently
porous to allow for chemical signaling between the co-cultured
cells. At the same time, the hydrogel channel wall should notallow the second injected hydrogel precursor component to diffuse into the vessel structures as this will cause clogging of theprinted structure. In this study, ﬂuorescence recovery after photobleaching (FRAP) served to investigate the permeability of the
2PP gelatin-based hydrogel on square pillar samples by using
FITC-Dextran of diﬀerent molecular weights as model molecules(Figure 6).
A decrease in diﬀusivity was observed as the molecular weightincreased up 250 kDa. For dextran molecules with a molecular© 2024 The Authors. Advanced Materials Technologies published by Wiley-VCH GmbH2365709x, 2024, 4, Downloaded from https://onlinelibrary.wiley.com/doi/10.1002/admt.202300718 by Uppsala University Library, Wiley Online Library on [22/05/2024]. See the Terms and Conditions (https://onlinelibrary.wiley.com/terms-and-conditions) on Wiley Online Library for rules of use; OA articles are governed by the applicable Creative Commons License

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Figure 5. 2PP printed structure for screening of the inner diameter after 10 days. Blue: 2PP ink. Red: Antonina-dextran. a) Top cross section of the 2PP
printed structure. b–c) Cross-section of the channels 30, 20, and 10 μm and 60, 40, and 20 μm, respectively. d) Inner diameter of the printed structure
after 1 and 10 days. e) Eccentricity of the printed structure after 1 and 10 days. Error bar from standard deviation (n = 3).

Figure 6. a) Diﬀusivity of 4, 40, 70, 250, and 500 kDa FITC-dextran molecular weights in the 2PP hydrogel after 24 h. (n = 6). b) Intensity proﬁle ofthe hydrogel (blue) and 250 kDa FITC-dextran (green) measured in the section highlighted within the red rectangle of image c). c) Confocal section ofa hydrogel sample immersed in the 250 kDa FITC-dextran solution. d) Schematic of the used woodpile conﬁguration of the 3D printing by 2PP. The
schematic shows two layers of the printing ﬁlaments of the printed sample displaying a crosslinking gradient from the core to the edge.

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www.advmattechnol.deweight of 500 kDa, no diﬀusion was observed inside the structures (Figure 6a). Consequently, we determined that growth factors produced by the encapsulated cells, at least up to 250 kDa,
will be able to diﬀuse from the cell-laden hydrogel to the endothelial cells during co-culture. In the speciﬁc case of collagenand ﬁbrin hydrogels, which have proteins with molecular weights
between 300–350 kDa, channel clogging could be expected. Although we cannot determine whether the low cut-oﬀ molecular weight for diﬀusion in the printed hydrogel is <300 kDa or
if the cross-linking process is so rapid that the hydrogel precursors do not diﬀuse across the channel walls, we did not observe any channel clogging in our experimental work. However,as a precaution, this fabrication strategy might not be suitablein cases where the second hydrogel has components in the precursor with molecular weights <250–300 kDa, as these might beable to diﬀuse through the 2PP channel wall and crosslink inside
the hollow structure, thus clogging the channel. Interestingly, weobserved that the 3D-printed structures did not display a uniform diﬀusion of the FITC-dextran molecules across the tested
sample (Figure 6b,c). In addition, the highest signal of FITCdextran molecules was observed in regions showing the lowest
hydrogel autoﬂuorescence signal (Figure 6b,c). Since brightﬁeld
images conﬁrmed the characteristic ﬁber-like structure of 2PP
printed structures (Figure S5, Supporting Information), the diffusion pattern was correlated to the voxel energy distributionthat presents lower energy at the voxel edges, causing an axial
crosslinking gradient in the ﬁber-like structure (Figure 6d).[95]The reduced crosslinking was associated with both higher porosity and diﬀusion as reported in previous studies.[96]2.1.5. The Fabrication Process Allows for both Straight andBranching Channels Perfusable for Week-Long Co-Cultures
To validate the compatibility of the proposed fabrication strategy for microphysiological studies, a co-culture of HUVECs and
HLFs was performed with both a straight channel and a more
complex vasculature model, (Figure S6, Supporting Information). The ﬁbroblast cells were chosen as they have been reported
as essential in the initial formation and maturation of vasculature systems.[92] Here, the HUVECs were seeded in the printed
channel after injection of the ﬁbroblast-laden ﬁbrin gel aroundthe 2PP structures. By turning the microﬂuidic chip upsidedown after ﬁbrin injection, ﬁbroblast sedimentation at the bottom of the device could be reduced and 3D cell growth enhanced.
Fibroblasts populated the whole ﬁbrin hydrogel and after 4–5
days of culture we could observe a higher HLF density close to
the HUVEC-lined microchannel, (Video S5, Supporting Information). From this work, we cannot directly conclude whether thisis due to HLFs migrating toward the vessels or a higher proliferation rate of the HLF in that area. However, both these events
are stimulated by a nutrient gradient from the perfused channels, suggesting that the 2PP hydrogel channel walls supported
the diﬀusion of nutrients as anticipated from the FRAP experiments. An optimal nutrient diﬀusion through the 2PP channel
was also hypothesized, as both cell types displayed viability >90%
after 7 days of culture (Figure 7f). The calcein staining used forthe live assay also displayed a higher ﬁbroblast concentration in

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proximity to the 2PP connectors and channel (Video S6, Supporting Information).
Of particular interest, we noted that the ﬁbroblast cells were
found in the space between the branching sections of the vessel
network, demonstrating a uniform spread of the ﬁbrin hydrogel
even around this complex structure. Around days 4–7, a retraction of the ﬁbrin gel from the glass substrate was observed. Fibrin retraction has been shown in previous studies.[97] However,the ﬁbrin retraction did not prevent ﬁbroblast growth around the2PP channel (Video S7, Supporting Information). Fibrin retraction was not observed in the section surrounded by the branched
2PP hydrogel of the structures (Figure 8b) and (Video S8, Supporting Information). We hypothesized that the branched 2PPhydrogel channels present a higher surface area compared to thesingle channel, thus promoting stability in the ﬁbrin hydrogel.
However, conﬁrmation of the hypothesis would require a moredetailed future study, outside the scope of this work.
The HUVECs’ initial distribution was higher at the two inlet
sides but the formation of a uniform cell layer across the wholechannel was observed after 2–6 days of culture under intermittent
ﬂow (Figure 7a–c) and (Video S9, Supporting Information).
Confocal microscopy imaging after 10 days of culture showeda conﬂuent HUVEC monolayer in the hydrogel channel(Figures 7d,e and 8b). The nuclei of the HUVECs inside thestraight hydrogel channel displayed a preferential orientation
along the channel axis, whereas cells cultured on the hydrogeldisks and glass of the microﬂuidic chip did not show preferentialalignment (Figure 7g). This compares well with previous studies
where the eﬀect of cell conﬁnement on cell orientation has beenstudied.[98,99] The HUVECs displayed F-actin-rich structures thatpenetrated the 2PP channel wall. Such structures were associated to ﬁlopodia, as involved in cell adhesion to ECM and guidance toward chemoattractants.[100] The presence of ﬁlopodia indicated the cells’ ability to penetrate the 2PP hydrogel (Figure 7d,e).
A cell ﬁlopodia depth gradient was observed from the channel
center to the inlets. The higher cell protrusion depth at the inlets could be associated with a longer cell-hydrogel interaction
time due to the initial seeding density distribution (Figure 7d).
HUVECs cultured in the absence of ﬁbroblasts did not display
F-actin protrusion penetration (Figure S7, Supporting Information). The results were in accordance with previous studies investigating the migration and sprouting of HUVEC co-culture withﬁbroblasts.[101]Finally, to demonstrate the formation of complex structureswith 2PP and the development of a co-culture platform with direct access of the two diﬀerent cells to each other, a 2PP hydrogel channel with cone holes along the sides was fabricated. A
truncated cone hole shape, with the small side of the cone cavity(10 μm) facing the external hydrogel, was chosen to create a highhydraulic resistance and prevent overﬂow of the ﬁbrin precursor
solution inside the 2PP channel. The aperture did not aﬀect themechanical stability of the 2PP hydrogel channel and no deformation was observed. The side holes created directional migration paths for the HUVECs toward the cell-laden ﬁbrin, resulting in HUVEC cell spreading on the outer surface of the 2PPhydrogel channel and spontaneous spouting into the ﬁbrin matrix (Figure 9; Video S10, Supporting Information). In the future,
we envision that this hybrid engineered-biologically driven system could provide a platform combining guided angiogenesis© 2024 The Authors. Advanced Materials Technologies published by Wiley-VCH GmbH2365709x, 2024, 4, Downloaded from https://onlinelibrary.wiley.com/doi/10.1002/admt.202300718 by Uppsala University Library, Wiley Online Library on [22/05/2024]. See the Terms and Conditions (https://onlinelibrary.wiley.com/terms-and-conditions) on Wiley Online Library for rules of use; OA articles are governed by the applicable Creative Commons License

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Figure 7. HUVEC growth is monitored inside the hydrogel channel. Green: the HUVEC cytoplasm a) cell distribution after seeding. b) cell distribution
after 24 h of culture. c) Cell distribution after 48 h of culture. d) After 10 days of culture, the hydrogel channel section (maximum projection of half heightchannel). The arrows indicate HUVEC ﬁlopodia. Yellow: nucleus, white: F-actin. e) cross-section of the hydrogel channel with HUVECs. Yellow: nucleus,
white: F-actin The arrows indicate HUVEC ﬁlopodia. f) Live-dead analysis for HUVECs and ﬁbroblasts after 7 days of culture. Error bar from standard
deviation (n = 3). g) Nucleus orientation distribution for HUVECs growth on the 2PP hydrogel disk, glass, and the hydrogel channel (n = 150).

and controlled vessel sprouting in a perfusable vessel-on-chipmodel.
The printed connectors at the 2PP ink/glass interface provided
a stable connection between the microﬂuidic chip and the printed
structure allowing perfusion of cell culture media. The shear
stress distribution for the chosen ﬂow rate around the two structures was calculated with numerical simulation by COMSOL toensure the chosen ﬂow rate exerted a shear stress that would
not be harmful toward the HUVEC cells, as shown in previous
studies[26,74] (Figure S8, Supporting information). The branchedstructure displayed a less uniform shear stress than the straightchannel due to the diﬀerent structural geometries.

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A key advantage of our strategy is that the microﬂuidic device
provides a platform where all necessary steps, 2-photon hydrogel
printing, second hydrogel casting, HUVEC seeding, cell media
perfusion, ﬁxation, staining, and high-resolution imaging of thecells, can be performed without the need to dismount or makeany changes to the platform, which signiﬁcantly simpliﬁes fabrication and handling of the sample.
The ability to create a perfusable vasculature model, combined with the ﬂexibility to guide cell invasion and interaction while not having constrains with cell encapsulation concentration in the hydrogel, opens up new possibilities for generating tissue models with physiological cell densities that© 2024 The Authors. Advanced Materials Technologies published by Wiley-VCH GmbH2365709x, 2024, 4, Downloaded from https://onlinelibrary.wiley.com/doi/10.1002/admt.202300718 by Uppsala University Library, Wiley Online Library on [22/05/2024]. See the Terms and Conditions (https://onlinelibrary.wiley.com/terms-and-conditions) on Wiley Online Library for rules of use; OA articles are governed by the applicable Creative Commons License

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Figure 8. Confocal images showing the vessel network with endothelial cells seeded inside and ﬁbroblasts in the surrounding ﬁbrin gel. a) Overview at
the mid height of the branched structure with HUVECs and ﬁbroblasts in co-culture. b) Magniﬁed section of the central branched structure (maximum
projection of half height channel, 25 μm). The HUVECs colonized the 2PP hydrogel channel generating a lumen while the ﬁbroblasts populated thevolume surrounding the hydrogel. c) and d) The cross section of 2 sections of the structure (indicated by a red dashed line).

3. Conclusion

Figure 9. HUVEC cells reach the ﬁbrin hydrogel passing through the created apertures in the 2PP structure. a) GFP channel image where the arrow indicates HUVEC sprouting. b) Brightﬁeld channel image where thedashed line indicates the contour of the inner 2PP hydrogel channel andthe shape of the cone-like holes.

In this work, we present a new fabrication strategy for making
a dual-hydrogel microvasculature model by combining the precision of 2PP and the cell compatibility of hydrogel casting. In
the presented two-step process, a vasculature network is ﬁrst deﬁned using 2PP inside a microﬂuidic device. Second, anothercell-laden hydrogel is injected around the 2PP-deﬁned structures.
The vasculature is ﬁnalized by seeding endothelial cells inside
the channel network. The study showed that this set-up supports
3D culture of ﬁbroblasts for up to 10 days in a perfusable vascularized hydrogel. The presented strategy not only preserves
the cell viability >90% of the 3D culture but also enables thegeneration of channels with feature sizes down to 10 μm, thus
replicating the capillary size of human tissue. The versatility of
2PP allowed the creation of side apertures in the 2PP channel
to promote HUVEC migration and sprouting, leading to a deﬁned angiogenesis density for more comparative studies in the
future.
By combining the reliable perfusion of the system with a new
high-resolution fabrication strategy, there is a unique opportunity to create innovative in vitro models of vascularized multihydrogel co-culture microphysiological systems.

4. Experimental Section
replicate vascularization, all within a portable microﬂuidic
device.

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Preparation of Multi-Hydrogel Structure: A commercially available 2PP
ink (U200, Bioinx) was used to print the sub-100 μm diameter channels.

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The 2PP ink was obtained by mixing 7.5 μl of the crosslinker solution(Dithiothreitol, provided as a ready-to-use solution by the supplier), 42.5 μl
of PBS (Sigma–Aldrich, concentration 1X) and 50 μl of the 2PP ink stock
solution heated to 37 °C (provided as a ready-to-use solution by the supplier) as instructed by the supplier. The hydrogel precursor solution was
vortexed and centrifuged (100g) to remove air bubbles and collect possible ﬁbers or debris at the bottom of the Eppendorf tube. Subsequently, thesolution was injected inside the microﬂuidic platform through one inletof the ﬂuidic device using a pipette. After the precursor solution injection,
the inlets and outlets were sealed with 3-mm punched PDMS disks (Super clear silicone sheet 0.5 μm, Silex Silicones). The hydrogel components
were mixed and injected in a sterile environment and all prints were performed within 3 h after mixing the hydrogel precursor components, as
instructed by the supplier.
The obtained 3D printed structures were surrounded by three diﬀerenthydrogels: ﬁbrin (Sigma–Aldrich), collagen (rat tail type 1 collagen, Gibco)
and a ﬁbrin–collagen mixture to display the versatility of the proposed approach. Fibrin and collagen were chosen since they have both been reported as suitable for 3D cell culture.[11] For ﬁbrin, a ﬁbrinogen solution
was mixed with thrombin (Sigma–Aldrich) in PBS to obtain a ﬁnal concentration of 2.5, 5, and 10 mg mL−1 ﬁbrinogen and 0.5 U mL−1 thrombin. For
collagen, a 3 mg mL−1 stock solution was diluted to a 2 mg mL−1 solution
by adding PBS and 1 m NaOH solution (11 μl of 1 m NaOH solution per
18 μl of 2 mg mL−1 collagen solution). For the ﬁbrin–collagen hydrogel, theﬁbrin and collagen stock solutions were mixed to obtain a 4.5 mg mL−1
ﬁbrin and 0.5 mg mL−1 collagen ﬁnal concentration. After mixing, all thehydrogel precursor solutions were immediately injected into the chip andincubated for 3 min to allow full crosslinking before immersing the wholechip in endothelial cell culture media.
For the 3D bioink biocompatibility assay, the protocol indicated by
the supplier was followed for the ﬁbroblast encapsulation. Brieﬂy, 50 μlof the 2PP ink stock solution was heated for 10 min at 37 °C before
adding 7.5 μl of the crosslinker solution and 42.5 μl of a PBS solution
with ﬁbroblasts to achieve a ﬁnal 2 × 106 cells mL−1 concentration. As
a control, ﬁbroblasts were also encapsulated in ﬁbrin hydrogels. The ﬁbrinogen stock solution was ﬁrst diluted in a PBS solution containing ﬁbroblasts and then mixed with a thrombin solution to achieve the ﬁnal
concentration of 5 mg mL−1 ﬁbrinogen, 0.5 U mL−1 of thrombin and2 × 106 cells mL−1 .
The ﬁbroblast-laden ﬁbrin for the multi-hydrogel structures were at aconcentration of 7.5 × 105 cells mL−1 , a cell density found to be suitable
via pilot experiments to prevent overgrowth of the ﬁbroblasts during the
10 days culture. After the ﬁbrin solution injection, the chip was rotatedevery 30 s for 3 min to prevent sedimentation of the cells at the bottom of
the device. The hydrogel was prevented from drying out by immersing thechip in cell media until further use.
Cell Culture: HUVEC-expressing green ﬂuorescent protein (GFP)(Angio-Proteomie, PELOBiotech GmbH) and HUVECs not expressing
GFP (ATCC) were cultured in a cell culture medium with an enhanced
growth kit provided by the supplier (Cellovations, Endothelial Cell Growth
Medium kit enhanced GFP, PELOBiotech). The culture ﬂasks were coatedto promote cell adhesion for 1 h (Speed Coating, PELOBiotech) prior toseeding cells that were used from passages 6 to 10.
Human Lung Fibroblasts (HLF) (Cellovations, PELOBiotech) were used
between passages 5 to 10 and cultured according to the supplier’s instructions (Cellovations Fibroblast Growth Medium kit enhanced, PELOBiotech). All cell types were kept in culture in an incubator at 37 °C with
5% CO2 and saturated humidity. The cell culture medium was changed
every 2 days.
Microﬂuidic Device Fabrication: Hydrogel analysis and cell cultures
were performed inside a PDMS-glass microﬂuidic device. The glass and
PDMS layers were air-plasma bonded (power: 200 W, time: 5 min, model:
Atto, Diener electronic GmbH) and then placed in contact at 100 °C for 2 h
to form a strong bond. Five diﬀerent designs were used for i) evaluatinghydrogel swelling, ii) performing FRAP experiments, iii) analyzing hydrogel cytocompatibility for 2D culture, iv) investigating cytocompatibility for
3D culture and v) combining two diﬀerent hydrogels. The more detailed

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layer-by-layer structures and 3D schematics of the microﬂuidic chips are
displayed in (Figures S9,S10, Supporting Information).
Device 1 (hydrogel swelling) consisted of a 250 μm-thick PDMS ring(Super clear silicone sheet 0.25 mm, Silex Silicones) with an inner diameter of 3 mm and outer diameter of 8 mm was bonded by air plasma to a
150-μm cover slip (VWR).
Device 2 (FRAP experiments and 2PP biocompatibility for encapsulated
ﬁbroblasts) consisted of a PDMS master (Figure S11, Supporting Information) was created with a 150 μm SU-8 photoresist laminate (DJ MicroLaminates, Inc.). PDMS prepolymer (Wacker Chemie AG) made with a 9:1
mixture ratio between base and curing agent was cast against the masterand degassed for 20 min before curing overnight in an oven at 60 °C. Afterremoving the PDMS structure, inlet and outlet holes were punched using
a 1 mm puncher. The bonding surfaces of the PDMS and glass slide were
plasma treated in air-plasma.
Device 3 (2PP ink biocompatibility and proliferation assay) consisted ofa 250 μm-thick PDMS layer (Super clear silicone sheet 0.25 μm, Silex Silicones) with three 1-mm punched holes sandwiched between the PDMS
chip and a cover slip. The device layers were manually aligned to ensure
overlapping of the channel and punched holes, followed by air-plasma
treatment.
Using Device 4 (biocompatibility for ﬁbroblast encapsulation), for chip
fabrication, the same protocol presented for device 2 was followed by achannel design that took advantage of hydrogel pinning to conﬁne the ﬁbrin hydrogel, as previously shown.[102]In Device 5 (multi-hydrogel constructs), the microﬂuidic chip consistedof a bottom channel (2 mm wide, 15 mm long and 0.25 mm high) and two
top channels (1 mm wide, 10 mm long and 0.2 mm high). The bottom
channel was fabricated by a cutter plotter (Craft ROBO Pro, Graphtec Corporation) using PDMS sheets (Super clear silicone sheet 0.25 μm, Silex
Silicones) while the two top channels were prepared by replica molding.
Five holes were punched in the PDMS layer obtained by replica molding.
The central hole served for the seeding of HUVECs while the 4 outer holes
were used for the injection of the ink, hydrogel or cell media perfusion. The
two microﬂuidic layers were separated by a 150 μm coverslip (Microscope
cover glasses, 24 mm × 24 mm, VWR) that included four holes. Two central holes allowed interconnection of the top and bottom channels, andtwo outer holes allowed for the injection of the ﬁbroblast encapsulatedhydrogel.
The coverslips in contact with the 2PP ink were functionalised withmethacrylated silane (Sigma–Aldrich) to improve adhesion of the hydrogel structures.[103] The treatment was initiated by an air-plasma clean(power: 200 W, time: 1 min, Model Atto, Diener electronic, Gmbh) followed
by immersion at room temperature in 2% w/w of (3-mercaptopropyl)
trimethoxysilane, 50% w/w ethanol, 47.7% w/w deionized water and 0.3%
w/w acetic acid for 20 min. Afterward, the chips were washed three times
with deionized water, dried under nitrogen gas and ﬁnally placed on a hotplate at 60 ˚C for 3 h. The functionalized chips were used within 1 monthfrom functionalization and protected from light during storage.
2PP Set-Up: All 3D printed structures were imported directly into
Think 3D software (UpNano, GmbH) as an STL ﬁle generated from Solidworks (Dassault Systèms). The objects were printed with the UpNano
One 3D printer (UpNano GmbH) in top–down mode with a 10x objective (0.4 NA, Olympus) by alternating layers scanned in the x and y directions (woodpile mode) to reduce asymmetric mechanical properties in the
structures. All 3D printing parameters were set in the Think3D software.
Prints were performed with a laser with a central wavelength of 790 nm and100 mW power. The refractive index for the 2PP ink hydrogel precursor solution was set as 1.36. All structures were printed with a hydrogel-glass
adhesion layer due to the oxygen crosslinking inhibition eﬀect near the
PDMS surface.[104,105]
Hydrogel Swelling: Swelling of the 2PP ink was assessed using a semiquantitative method employed in previous studies.[53,106] First, 25 cubes(100 μm × 100 μm × 100 μm) were printed (Figure S12, Supporting Information). For the retained cubes with little distortion, the size of the top
structure surface (i.e., the surface furthest away from the hydrogel-glass
interface) was measured via confocal microscopy after 24 h incubation in© 2024 The Authors. Advanced Materials Technologies published by Wiley-VCH GmbH2365709x, 2024, 4, Downloaded from https://onlinelibrary.wiley.com/doi/10.1002/admt.202300718 by Uppsala University Library, Wiley Online Library on [22/05/2024]. See the Terms and Conditions (https://onlinelibrary.wiley.com/terms-and-conditions) on Wiley Online Library for rules of use; OA articles are governed by the applicable Creative Commons License

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www.advmattechnol.decell media. Then, from the surface area, the side length of each top surface
was calculated. The degree of swelling was determined by comparing thedimensions of the square side to the dimension in the original CAD ﬁle,according to the following formula:
Swelling (%) =SCUB − SCAD
SCAD

(1)

where SCUB is the top side length of the cube and SCAD is the top side
length of the CAD ﬁle.
Diﬀerent z intervals (1–9 μm) and scanning speeds (200–
1,000 mm s−1 ) were screened for printing at a constant laser power(100 mW). The space between two adjacent samples was chosen to
avoid printed stitching lines from overlapping in the samples. For the
sample displaying the least swelling, the hydrogel-glass interface wasalso measured via confocal microscopy to investigate the printing quality
when not aﬀected by hydrogel swelling.
Single and Multi-Hydrogel Structure Fabrication: For the multi-hydrogel
structures, the 2PP channel and connector wall thicknesses were set to 40
and 175 μm, respectively. After printing in device ﬁve, the non-crosslinked
hydrogel was gently removed with warm media. The ﬁbrin, collagen, andﬁbrin–collagen precursor solutions were injected around the printed channels by manually pipetting (Figure S13, Supporting Information). The
single-hydrogel structure was printed with 200 μm-thick walls for both thechannel and the connectors. Overnight incubation in cell media at 37 °C
allowed the hydrogel structures to equilibrate before being evaluated.
Fluorescence of the hydrogel allowed to image its structure via a confocal microscope (Leica-SP8, 10× objective and 0.3 NA). Reproducibility ofthe printed structures was evaluated with x-y confocal images, and the inner diameter of the hydrogel channel was compared with the dimensions
in the CAD design. In addition, images were recorded in reﬂection mode
to visualize the ﬁbrin, collagen and ﬁbrin–collagen network surrounding the 2PP structure and evaluate the eﬀect of the non-crosslinked ink
removal.
For the investigation of minimum achievable feature sizes, ﬁve different hollow structures were printed with inner diameters of either 10,
20, 30, 40, or 60 μm and a constant wall thickness of 40 μm. After removal of the non-crosslinked hydrogel ink, a 500 kDa Antonina solution(1 mg mL−1 , Sigma–Aldrich) was injected inside the vessel structure atday 1 and 10 to measure the diameter and eccentricity of the channels.
The images were taken by confocal microscopy (Leica, SP8, 10x air objective 0.3 NA) with a z-stack of 200 μm and analyzed with CellProﬁler
software.[107]
FRAP Experiments: FITC-dextran (Sigma–Aldrich) with ﬁve molecularsizes, 4, 40, 70, 250, or 500 kDa (concentration 50 μM, Sigma) were used.
The hydrogel precursor solution was manually injected with a pipette into
the microﬂuidic device and 6 structures (300 μm × 300 μm × 150 μm) wereprinted at the bottom of the channel (Figure S14, Supporting Information).
The chip was then immersed in cell media overnight to remove the noncrosslinked ink. Afterward, the hydrogel samples were incubated in FITCdextran solution for 24 h to reach equilibrium. FRAP was then monitored(recorded time 60 s, frame interval 0.5 s) with an inverted confocal microscope (Leica, SP8, 25x water objective, 0.95 NA). For each FITC-dextran
molecular size, six sample recovery sequences were processed with the“FRAP analysis” library from Matlab.[108]The diﬀusion pattern observed in the hydrogel structures was analyzed
with Fiji by plotting the intensity proﬁle of the hydrogel and FITC-dextran
solution.[109]2PP Bioink and Cell Interaction: For 2D culture assessment, a disk witha 1 mm diameter and 100 μm thickness was printed at the bottom of thewells on device three. The non-crosslinked 2PP ink was gently removed
by injecting fresh media into the microﬂuidic chip, and the printed structures kept in an incubator overnight to equilibrate. Non-GFP-expressing
HUVECs and GFP-expressing HUVECs were seeded in the microﬂuidic
chip at a concentration of 104 cells cm−2 for cell viability and proliferation,
respectively. Cell viability was measured at 24 h after seeding (n = 5). Proliferation was monitored over a period of 10 days with measurements collected at days 1, 3, 7, and 10 (n = 9). As control, the same microﬂuidic chip

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design was treated with an adhesion-promoting solution recommended by
the cell provider (SpeedCoating, PELOBiotech) and cells were cultured on
this surface. For 3D culture biocompatibility, the ﬁbroblast-laden 2PP ink(2 × 106 cells mL−1 ) was injected in device two to fabricate cube structures(500 × 500 × 250 μm3 ) with the optimized printing parameters. After printing, the non-crosslinked hydrogel was removed by injecting warm media.
For the control experiment, the ﬁbroblast-laden ﬁbrin precursor solution
was injected into the central channel of device four and kept in the incubator for 3 min before adding cell media on the side channels. Cell viabilityof HUVECs and ﬁbroblasts was tested 24 h after hydrogel crosslinking byperforming a live/dead assay (Thermo Fisher Scientiﬁc) with 0.75 μl mL−1
propidium iodide and 2 μl mL−1 calcein AM, with concentrations from thestock solution provided by the supplier. The samples were incubated for
15 min after assay solution injection and then washed three times with
PBS before imaging by confocal microscope (Leica SP8 10 × 0.4 NA air
objective). Cell proliferation was measured by total nuclei counts and total cell coverage area. On day 10, cells were stained with a live nucleus
ﬂuorescent dye (Spy DNA 555, Spirochrome) by direct injection inside themicroﬂuidic chip 1 h before images were captured. The same live/deadprotocol was followed for assessing the viability of ﬁbroblast and HUVEC
cell viability in the multi-hydrogel structure. Image analysis for both 2D
and 3D cultures were performed using CellProﬁler.
Shear Stress Simulation: The wall shear stress experienced by the cells
in the diﬀerent hydrogel channel designs was determined with COMSOL Multiphysics software (Version 6.0, COMSOL Inc). Hydrogel channel
structures were designed in Solidworks software (version 2021). After importing the structure into the COMSOL software, the assumptions usedin the models were Newtonian and incompressible ﬂuid with the dynamic
viscosity of pure water. In addition, the following settings were used: The
no-slip boundary condition ensured zero velocity at the wall, and the atmosphere pressure was set at the outlet. Inlet boundary conditions served toinvestigate the shear stress for the applied ﬂow. The simulation was performed with the laminar ﬂow module and ﬁner mesh. The ﬂow-induced
deformation of the hydrogel channel was assumed to be negligible after
no deformation was observed in the channel during perfusion by microscope imaging (Lumascope 560, Etaluma).
Perfusable Multi-Hydrogel Construct for Co-Culture of HUVECs and Human Lung Fibroblasts: After 2PP hydrogel channel printing, a ﬁbroblastladen ﬁbrin solution (7.5 × 105 cells mL−1 ) was manually injected witha 100 μl pipette tip to surround the vessel structures, and incubatedovernight. The next day, HUVECs (1.5 × 107 cells mL−1 ) were seeded
inside the printed network by pipette injection. This cell concentration
was chosen experimentally to ensure a suﬃcient number of cells in thehydrogel channel. After 12 h, the channels were connected to a syringe
pump (Nemesys, low pressure module) so that the hydrogel channel media could be refreshed with an intermittent ﬂow (ﬂow rate: 0.1 μl min−1 ,shear stress: 0.6 dyne cm−2 , 3 min perfusion every 4 h) for 10 days to promote endothelial monolayer formation. The cell culture was monitoredwith an incubator microscope (Lumascope 560, Etaluma) and the use of acustomized microﬂuidic carrier allowed user-friendly handling, as shownin previous studies[110,111] (Figure S15, Supporting Information).
At day 10, the samples were ﬁxed with 2% paraformaldehyde for
15 min. (Thermo Fisher Scientiﬁc) and stained for F-actin (SPY620 actin,
Spirochrome) and nuclei (SPY555 DNA, Spirochrome), before being imaged with a confocal microscope (Leica SP8, 25X water objective, 0.95
N.A).
Cell Nucleus Orientation: At day 7, the cells cultured on the hydrogeldisks (cell seeding and culture protocol are shown in Section 4.3.6), inside
the microﬂuidic chip channels and on the hydrogel channels (cell seeding
and culture protocol are shown in Section 4.3.8) were stained (SPY555
DNA, Spirochrome) and imaged by confocal microscopy (Leica SP8). For
the hydrogel disks, three samples in diﬀerent microﬂuidic chips were imaged. Regarding the microﬂuidic chip channel, a surface area of 1 mm2
was acquired. Finally, for the hydrogel channel, the section between thehydrogel connector was imaged. Then, the nucleus orientation of all theconditions was measured with CellProﬁler.
Statistical Analysis: For the nuclei orientation, at least 150 nuclei wereanalysed per condition. As the alignment of the nuclei was always centred© 2024 The Authors. Advanced Materials Technologies published by Wiley-VCH GmbH2365709x, 2024, 4, Downloaded from https://onlinelibrary.wiley.com/doi/10.1002/admt.202300718 by Uppsala University Library, Wiley Online Library on [22/05/2024]. See the Terms and Conditions (https://onlinelibrary.wiley.com/terms-and-conditions) on Wiley Online Library for rules of use; OA articles are governed by the applicable Creative Commons License

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www.advmattechnol.de

on 0˚ there was no diﬀerence between the means of this study. Instead,they diﬀer because they had diﬀerent variances. For the nucleus orientation experiment RStudio was therefore statistically tested if there was adiﬀerence in variance between the diﬀerent conditions using an Ansari–
Bradley test with a conﬁdence level of 95%.
For hydrogel swelling, there was a diﬀerence in means between the conditions and thus a more standard statistical approach was applied. The
variable swelling was transformed according to the Box-Cox test to meet
the assumption of normality. Diﬀerences between factor variables were
determined using a Tukey post hoc test with a conﬁdence level of 95%.
P-values <0.05 were considered statistically signiﬁcant as follows where* denotes p < 0.05, ** denotes p < 0.01 and *** denotes p < 0.001.

Supporting Information
Supporting Information is available from the Wiley Online Library or fromthe author.

Acknowledgements
This research was funded by the European Research Council (ERC) under the European Union’s Horizon 2020 research and innovation program(grant agreement No 757444) and the Knut and Alice Wallenberg Foundation (grant number WAF 2016.0112). We acknowledge Myfab Uppsala forproviding facilities and experimental support, especially Milena Moreirafor the support with the 2-photon printer. Myfab is funded by the Swedish
Research Council (2019-00207) as a national research infrastructure. We
also greatly acknowledge the eﬀorts by Susan Peacock and Samuel Lindner, Uppsala University, for the careful proof-reading of the manuscript
and the fruitful discussion about the ﬁbrin hydrogel and cell culture, respectively. Finally, we acknowledge Elsa Lagerquist, SLU, and Estefania
Echeverri, Uppsala University, for the valuable help with the statistical analysis and the kind donation of collagen, respectively.

Conﬂict of Interest
The authors declare no conﬂict of interest.

Author contributions
F.C., L.B., and M.T. conceptualized the study, F.C. and L.B. conceived andoptimized the 2-step hydrogel fabrication process and the device designs,
F.C., L.B., H.P., and M.T. planned the experiments, F.C performed the experiments and collected the data, F.C., L.B., and H.P. analyzed the data.
F.C. wrote the original draft. F.C., L.B., H.P., and M.T. wrote, reviewed,
and edited the manuscript. L.B. and M.T. supervised the research. M.T. acquired the funding. All authors edited and approved the ﬁnal manuscript.

Data Availability Statement
The data that support the ﬁndings of this study are available from the corresponding author upon reasonable request.

Keywords2-photon polymerization 3D printing, microphysiological systems, multihydrogel structure, organ-on-chip, vasculature model
Received: May 8, 2023
Revised: November 15, 2023
Published online: January 4, 2024

Adv. Mater. Technol. 2024, 9, 2300718

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