The authors thank Dr. Jeff Caplan and Mr. Michael Moore at the Delaware Biotechnology Institute Bio-Imaging Center for their support with confocal microscopy. Access to microscopy equipment was supported by the National Institutes of Health (NIH) shared instrumentation grants (S10 RR0272773 and S10 OD016361) and the State of Delaware Federal Research and Development Grant Program (16A00471). This research was supported by grants from the Institutional Development Award (IDeA) from the NIH National Institute of General Medical Sciences (P20GM103446), the American Cancer Society (14-251-07-IRG), the University of Delaware Research Foundation (14A00778), the Cancer Prevention and Research Institute of Texas (CPRIT) (RR140013), and the NIH National Library of Medicine (4 R00 LM011390-02).

<ol>
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The authors have nothing to disclose.

Critical in overriding the standard functions of the microscope software to enable the use of virtual masks for image-guided, laser-based degradation, the microscope macro must be setup and manipulated carefully. How the microscope software interfaces with the microscope macro can sometimes be counterintuitive, and much effort was invested in determining the optimal settings in both programs to develop this method. A general understanding of basic laser-scanning confocal microscope use is recommended, especially with the &#34;Stage&#34; and &#34;Focus&#34; windows for finding and correctly positioning the hydrogel in the x, y, and z dimensions, before attempting laser-based degradation. Additionally, the fabrication of an inlet channel is critical to the protocol; suspended fluorescent species must be able to enter the microfluidic systems for successful imaging and network characterization. It is important to note that this protocol applies to a specific laser-scanning confocal microscope configured with a specific femtosecond pulsed laser, running specific versions of the microscope software and the microscope macro (see details in the List of Specific Materials/Equipment). We have discovered that newer versions of the microscope macro lack the necessary control and functionality needed for image-guided laser control. While other microscope platforms and pulsed laser sources can be used, this protocol applies specifically to this system and will need modification according to the platform, laser source, and software implemented. The underlying principles throughout this protocol still apply, however.
A potential modification to this protocol involves changing the hydrogel composition. Here, we utilized 5 wt%, 3.4 kDa PEGDA hydrogels, but laser-based degradation (for purposes aside from microfluidic network generation) has been implemented to degrade both synthetic and natural hydrogels, including PEGylated fibrinogen21,22, silk protein hydrogels23, and collagen24. Adjustment of the laser power, scan speed, spacing between Z-slices, and number of repetitions will aid in determining the optimal parameters to fabricate microfluidic networks in other hydrogel formulations.
One current limitation of the technique is the overall volume of hydrogel that can be degraded in a feasible amount of time. To create open voids or microfluidic features within the hydrogel, the laser dwell time must be at or above 8.96 &#181;s/pixel (or a laser scan speed of 0.021 &#181;m/&#181;s) when using a laser fluence of 37.7 nJ/&#181;m2. With these settings, it takes 1.4 h to degrade vessels within a 0.014 mm3 volume (as seen in Figure 1). Using a laser dwell time of 4.48 &#181;s/pixel or below, the energy delivered is not enough for full degradation of the hydrogel formulation used here. Implementing a different hydrogel composition could overcome this limitation. The use of photolabile gels that contain light-sensitive components34-36 or hydrogels that have large multiphoton cross-sections21-24 are good options that would enable the use of less energy and would result in faster degradation. With respect to dimensional limitations, features have been fabricated up to 1.5 mm deep in the hydrogel7. The depth achievable is a function of the working distance of the objective and can be increased using ultra-long working distance objectives optimized for in vivo imaging. The smallest measured channel7 created using a 20X (NA1.0) water immersion objective had a width of 3.3 &#181;m and depth of 8.9 &#181;m, which is on par with the size of the smallest channels generated in Figure 1. While other labs have used lower NA objectives21,23,24, we anticipate that microfluidic networks with smaller features can be generated using higher NA objectives at the expense of a reduced working distance. Ultimately, however, the resolution of the technique is a function of the point spread function of the focused laser beam, the laser scanning properties, the amount of energy delivered by the laser, and the laser absorption properties of the material being degraded.
Furthermore, laser properties (speed, fluence, spacing between virtual masks, and number of repetitions) must be optimized based on hydrogel properties (crosslink density, macromer/monomer molecular weight, weight percent, and type of hydrogel: protein-based vs. synthetic), as these will inherently alter interactions with the laser and thus the ultimate resolution of the process. As the energy delivered is also influenced by the objective used, additional optimization is required when switching between objectives. With respect to the costs involved, access to a microscope with a pulsed laser is required, and while many different objectives can be used, those with a high NA and long working distance (for in vivo imaging) can be expensive.
Above and beyond the protocol detailed here, microfluidic networks generated using this technique can also be functionalized with cell-adhesive peptides post-channel fabrication to induce endothelial cell adhesion and lumen formation7. Additionally, the incorporation of cell-degradable peptide sequences in the bulk material9 prior to channel degradation could allow for the study of cell migration into and through the hydrogel. For continuous fluidization of the microfluidic network within the hydrogel, hydrogels can be photopolymerized in larger fluidic devices or housings, as demonstrated in Figures 2 and 37.
The generation of vascular networks via vasculogenic or angiogenic self-assembly8-12 provides a straightforward approach to induce vascularization throughout a relatively large hydrogel volume. While this approach results in perfusable fluidic networks, it is difficult to directly control the size, tortuosity, density, and overall network architecture. Due to this limitation, the flow profile and shear rates in the vasculature may differ between experiments. Alternatively, microfabrication techniques13-16 allow for direct control over network architecture but are often limited by their inability to generate small, capillary-sized channels or the fabrication of dense networks that mimic in vivo vascular architecture. While the laser-based hydrogel degradation technique outlined here has been newly repurposed for microfluidic generation, it simultaneously overcomes both the architectural control and size-based limitations of existing microfabrication methods by enabling the creation of 3D biomimetic microfluidic networks that recapitulate the density, tortuosity, size range, and overall architecture of in vivo vasculature. Furthermore, multiple fluidic networks can be generated in a single hydrogel, allowing the study of inter-network transport7. For tissue engineering applications that strive to more closely replicate in vivo transport processes in vitro, the highly resolved hydrogel-embedded microfluidic networks outlined here are well suited. We anticipate that this protocol will be useful in developing tissue constructs that more accurately mimic in vivo transport for use as drug screening devices and in vitro disease models.

The vascular system, consisting of both lymphatic and cardiovasculature, forms highly dense networks that are essential for the transport of nutrients and oxygen and for the removal of metabolic waste. Accordingly, cells residing in vascularized tissues are never more than 50-100 &#181;m away from a vessel1. The ability to reproduce in vivo vascular architecture in vitro is critical to accurately model in vivo transport processes using engineered constructs. With a recent drive to develop organ-on-a-chip devices2 for high-throughput drug screening3,4 and disease modeling5,6, methods to create microfluidic networks that recapitulate in vivo-like transport in synthetic or natural hydrogels are drawing significant interest. To enhance the biomimicry of microtissues used in these devices, we developed an image-guided, laser-based hydrogel degradation method that utilizes three-dimensional (3D) image stacks of native vasculature as templates to generate vascular-derived microfluidic networks embedded in PEGDA hydrogels7. This protocol outlines the use of a commercially available laser-scanning confocal microscope equipped with a femtosecond pulsed laser to fabricate vascular-derived, biomimetic microfluidic networks in PEGDA hydrogels via image-guided, laser-based degradation.
Current approaches to fluidize hydrogels include the induction of vasculogenic8-10 or angiogenic10-12 endothelial cell self-assembly and microfabrication techniques to create pre-defined channels for endothelialization13-16. While self-assembled networks recapitulate the density and complex architecture of microvasculature, they are often more permeable than in vivo networks11,17,18, which may be problematic in modeling transport for drug screening applications. Self-assembled networks consist of physiologically relevant, capillary-sized vessels, but they can be difficult to integrate with bulk fluid flow due to limitations in generating larger arteriole-sized vessels. As there is no direct control over the assembly of these networks, the final architecture can vary from sample to sample, making it difficult to repeatedly produce networks with the same fluid flow and transport properties.
To generate 3D, hydrogel-embedded microfluidic networks with repeatable geometry and well-defined architecture, a number of microfabrication techniques have been developed, including modular assembly13, 3D printing of sacrificial materials16, direct-write assembly14, and omnidirectional printing15. Applying these methods, microfluidic architecture, and therefore fluid flow and transport properties, can be repeatedly fabricated across many constructs. A major limitation of these approaches, however, is the inability to create microfluidic networks with capillary-sized features, 4 to 10 &#181;m19. Most microfabrication techniques are often limited to features ranging from 150 to 650 &#181;m in diameter13,16. Some existing techniques are capable of generating hierarchical networks with channels across a wide diameter range, 10 to 300 &#181;m for direct-write assembly14, and 18 to 600 &#181;m for omnidirectional printing15, but they are limited in their ability to generate dense networks or to produce multiple microfluidic networks in close proximity within a single construct7.
To overcome some of these limitations, we developed an image-guided, laser-based hydrogel degradation technique that allows repeatable fabrication of biomimetic, hierarchical microfluidic networks that recapitulate the architecture of in vivo microvasculature. To do so, a 790 nm, 140 femtosecond (fs) pulsed laser operating at 80 MHz is raster-scanned in desired 3D locations within a hydrogel, as defined by images of in vivo vasculature. We speculate that the degradation process operates through the laser-induced optical breakdown of water, the resultant plasma formation, the subsequent rapid thermoelastic expansion of water, and the local degradation of the hydrogel as the water expands20. This mechanism varies slightly from laser-based degradation of protein-based hydrogels21-24. Unlike PEGDA, which has a low multiphoton cross-section, proteins often have a large multiphoton cross-section and are therefore degraded via a multiphoton absorption-induced chemical breakage23. To generate image-guided microfluidic networks, the laser shutter is controlled by image-derived virtual masks, which consist of a mosaic of regions of interest9 that define the microfluidic architecture. Using this approach, we have demonstrated the ability to fabricate 3D vascular-derived biomimetic microfluidic networks, which recapitulate the dense and tortuous architecture of in vivo vasculature, in order to locally control the porosity of hydrogels by altering the amount of energy delivered during degradation. We have also been able to generate two independent microfluidic networks that intertwine in close proximity (15 &#181;m) but never directly connect7. We have also demonstrated the ability to endothelialize laser-degraded microchannels through post degradation functionalization with the integrin ligating peptide sequence, Arginine-Glycine-Aspartic Acid-Serine (RGDS), to promote endothelial cell adhesion and lumen formation7.
With this protocol, the generation of complex microfluidic networks in PEGDA hydrogels is made possible via image-guided, laser-based degradation using a commercially available microscope accessible on many university campuses. As the degradation process is guided by digital, virtual masks, this fabrication technique is amenable for the creative design of microfluidic networks, allowing its use in a wide variety of applications. We anticipate that the method described here will be most advantageous in designing biomimetic organ- and human-on-a-chip devices capable of replicating biological transport processes important in modeling drug transport2. This fabrication technique may also be of interest for the generation of in vitro disease models, including cancer metastasis5,6 and blood-brain barrier models25. As laser-based hydrogel degradation has previously been used to create tracks for the guidance of neuronal outgrowth21-23, the image-guided extension of this technique could prove useful in advanced tissue engineering strategies to position cells in user-defined 3D spatial arrangements.

<table><tbody><tr><td>MATLAB</td><td>Mathworks</td><td>R2015a</td><td>named &quot;programming software&quot; in protocol; refer to source 9 for details on algorithm</td></tr><tr><td>FIJI (Fiji is Just Image J)</td><td>NIH</td><td>version 1.51a</td><td>named &quot;image processing software&quot; in protocol</td></tr><tr><td>LSM 780 Confocal Microscope</td><td>Zeiss</td><td></td><td>named &quot;laser-scanning confocal microscope&quot; in protocol; for laser-based hydrogel degradation</td></tr><tr><td>Zen 2010B SP1</td><td>Zeiss</td><td>release version 6.0</td><td>named &quot;microscope software&quot; in protocol; for use on Zeiss LSM-780</td></tr><tr><td>Multitime, 2010</td><td>Zeiss</td><td>v16.0</td><td>named &quot;microscope macro&quot; in protocol; for use on Zeiss LSM-780 (in conjunction with Zen)</td></tr><tr><td>Objective W Plan-Apochromat 20X/1.0 DIC D=0.17 M27 75 mm</td><td>Zeiss</td><td>421452-9880-000</td><td>for use on Zeiss LSM-780</td></tr><tr><td>Chameleon Vision II Modelocked Ti:S Laser</td><td>Coherent</td><td></td><td>named &quot;high-powered pulsed laser&quot; in protocol</td></tr><tr><td>Sodium chloride</td><td>Sigma-Aldrich</td><td>S5886-1KG</td><td></td></tr><tr><td>HEPES</td><td>Sigma-Aldrich</td><td>H3375-250G</td><td></td></tr><tr><td>Triethanolamine (TEOA)</td><td>Sigma-Aldrich</td><td>90279-100ML</td><td>flammable; skin and eye irritant; work with in a fume hood</td></tr><tr><td>Sodium hydroxide</td><td>Sigma-Aldrich</td><td>S5881-500G</td><td></td></tr><tr><td>150 mL Vacuum Filtration Cups with 0.2 &amp;micro;m PES Membrane</td><td>VWR</td><td>10040-460</td><td></td></tr><tr><td>PEGDA</td><td>synthesized in house</td><td></td><td>refer to source 33 for synthesis methods; store under argon</td></tr><tr><td>Eosin Y disodium salt</td><td>Sigma-Aldrich</td><td>E6003-25G</td><td></td></tr><tr><td>1-vinyl-2-pyrrolidinone (NVP)</td><td>Sigma-Aldrich</td><td>V3409-5G</td><td>store under argon; carcinogenic; work with in a fume hood</td></tr><tr><td>3M Double Coated Tape, 9500PC, 6.0 mil</td><td>Thomas Goldkamp</td><td>37728</td><td></td></tr><tr><td>Flexmark 90 PFW Liner</td><td>FLEXcon</td><td>FLX000620</td><td>backing for handling of double coated tape</td></tr><tr><td>Model SC Plotter (adhesive cutter)</td><td>USCutter</td><td>SC631E</td><td>used to cut adhesive in ring shapes to connect coverslips to petri dishes</td></tr><tr><td>60 mm Petri Dish with 20 mm Hole</td><td>MatTek Corporation</td><td>P60-20-F-NON</td><td></td></tr><tr><td>High Intensity Illuminator (white light source)</td><td>Fiber-Lite</td><td>4715MS-12WB10</td><td></td></tr><tr><td>Power Meter</td><td>Newport</td><td>1916-R</td><td>detect power at 524 nm when using white light source</td></tr><tr><td>Slim Profile Wand Detector</td><td>Newport</td><td>918D-ST</td><td>for use with power meter</td></tr><tr><td>Sylgard 184 Silicone Elastomer Kit</td><td>Dow Corning</td><td>3097358-1004</td><td>used to make PDMS molds; refer to source 7 for methods</td></tr><tr><td>TMPSA-Functionalized #1.5 Coverslips, 40 mm Round</td><td>synthesized in house</td><td></td><td>refer to source 7 for methods</td></tr><tr><td>Dextran, Fluorescein, 2,000,000 MW, Anionic, Lysine Fixable</td><td>Life Technologies</td><td>D7137</td><td>can use alternative tagged dextrans; 2,000 kDa does not diffuse readily into a 5% 3.4 kDa PEGDA hydrogel</td></tr><tr><td>1 mL Syringe, Luer-Lok</td><td>BD</td><td>309628</td><td></td></tr><tr><td>Acrodisc Syring Filter, 0.2 &amp;micro;m Supor Membrane, Low Protein Binding</td><td>Pall</td><td>PN 4602</td><td></td></tr><tr><td>sticky-Slide VI&lt;sup&gt;0.4&amp;nbsp;&lt;/sup&gt;</td><td>Ibidi</td><td>80601</td><td>microfluidic devices that can be used to house hydrogels</td></tr></tbody></table>

image-guided, laser-based fabrication of vascular-derived microfluidic networks

This detailed protocol outlines the implementation of image-guided, laser-based hydrogel degradation for the fabrication of vascular-derived microfluidic networks embedded in PEGDA hydrogels. Here, we describe the creation of virtual masks that allow for image-guided laser control; the photopolymerization of a micromolded PEGDA hydrogel, suitable for microfluidic network fabrication and pressure head-driven flow; the setup and use of a commercially available laser scanning confocal microscope paired with a femtosecond pulsed Ti:S laser to induce hydrogel degradation; and the imaging of fabricated microfluidic networks using fluorescent species and confocal microscopy. Much of the protocol is focused on the proper setup and implementation of the microscope software and microscope macro, as these are crucial steps in using a commercial microscope for microfluidic fabrication purposes that contain a number of intricacies. The image-guided component of this technique allows for the implementation of 3D image stacks or user-generated 3D models, thereby allowing for creative microfluidic design and for the fabrication of complex microfluidic systems of virtually any configuration. With an expected impact in tissue engineering, the methods outlined in this protocol could aid in the fabrication of advanced biomimetic microtissue constructs for organ- and human-on-a-chip devices. By mimicking the complex architecture, tortuosity, size, and density of in vivo vasculature, essential biological transport processes can be replicated in these constructs, leading to more accurate in vitro modeling of drug pharmacokinetics and disease.

To demonstrate the implementation of image-guided, laser-based hydrogel degradation to generate a 3D, vascular-derived microfluidic network, we utilized a 3D image stack of mouse brain vasculature that was acquired via knife-edged scanning microscopy (KESM)26,27. A selected 3D region of the larger microvascular dataset was converted into a series of virtual masks for image-guided microfluidic network formation, as described above. After fabrication, the resulting microfluidic network was perfused with a solution of FITC-labeled, 2,000 kDa dextran and imaged via confocal microscopy. The image stack of the in vivo vasculature that defined the network architecture (Figure 1A and C) and the fabricated microfluidic network (Figure 1B and D) were used to generate Z-projections (Figure 1A and B) and 3D renderings (Figure 1C and D). The comparison of the images demonstrates that image-guided, laser-based hydrogel degradation enables the fabrication of 3D biomimetic microfluidic networks that recapitulate the size, tortuosity, and complex architecture of in vivo vasculature. The technique is amenable to fabrication of networks containing a wide range of diameters; within Figure 1, channels ranging from 3.3 to 28.8 &#181;m in diameter were generated. Note that the larger rectangular structure in the top left of Figure 1B and in the left corner of Figure 1D is the inlet channel to introduce flow into the network.
<img alt="Figure 1" src="/files/ftp_upload/55101/55101fig1.jpg" /> 
Figure 1: Vascular-Derived Microfluidic Network. A series of virtual masks, derived from an image stack of mouse brain vasculature (A and C), were used to fabricate a 3D biomimetic microfluidic network embedded in a PEGDA hydrogel (B and D) via image-guided, laser-based degradation. The microfluidic network was perfused with 2,000 kDa FITC-labeled dextran and imaged via confocal microscopy. Z-projections (A and B) and 3D renderings (C and D) of the two networks demonstrate the ability to generate microfluidic networks that recapitulate the density, organization, and overall architecture of in vivo vasculature. The arrow (D) marks the rectangular inlet created to introduce the dextran. The scale bar represents 50 &#181;m. <a href="https://cloudflare-jove-com.bdigitaluss.remotexs.co/files/ftp_upload/55101/55101fig1large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
We have implemented two methods, pressure heads and syringe pumps, to initiate fluid flow in these embedded microfluidic networks. For example, a 30 by 30 &#181;m rectangular channel was fabricated to connect two wells in a micromolded PEGDA hydrogel (Figure 2A), with fluid flow driven via a pressure head7. Generated in the left well, the pressure head induced flow through the microchannel and into the well on the right. In Figure 2A, an initial front of tetramethylrhodamine (TRITC)-labeled, 70 kDa dextran was observed moving through the channel using time-lapse microscopy. Later, in Figure 2B, a 10-&#181;m diameter fluorescent polystyrene sphere flowed from the left well, through the channel, to the well on the right. With the duration of flow controlled by the volume of fluid present in the micromolded well at the inlet, and the flow-rate controlled by the hydrostatic gradient generated between the inlet and outlet well, pressure head-driven flow in micromolded hydrogels provides a simple method for flow induction, without the need for an external housing or a syringe pump.
<img alt="Figure 2" src="/files/ftp_upload/55101/55101fig2.jpg" /> 
Figure 2: Pressure Head-Driven Flow in a Micromolded PEGDA Hydrogel. PEGDA was micromolded to form a hydrogel containing two wells connected by an embedded microchannel. A pressure head was generated in the left well to initiate flow of TRITC-labeled, 70 kDa dextran (A1-A3) and a 10-&#181;m polystyrene sphere (B1-B3) through a 30 by 30 &#181;m rectangular channel from one well to the other. The scale bar represents 50 &#181;m. <a href="https://cloudflare-jove-com.bdigitaluss.remotexs.co/files/ftp_upload/55101/55101fig2large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
Alternatively, to generate constant fluid flow within microfluidic networks, syringe pump-driven flow can be initiated by photopolymerizing the hydrogel inside of a secondary microfluidic device with inlet and outlet ports for syringe pump interfacing. In Figure 3A, a commercially available, pre-fabricated microfluidic device is shown with a 1-mm band of hydrogel photopolymerized across the width of the device7. Laser-based degradation was implemented to fabricate microfluidic channels and networks in housed hydrogels using the same protocol described here for freestanding hydrogels. Syringe pump-generated flow can be seen in Figure 3B, where a 20-&#181;m diameter cylindrical channel was fabricated in a hydrogel housed in a microfluidic device. In this 20-&#181;m diameter channel, individual 2-&#181;m fluorescent polystyrene spheres are easily resolved without fluid flow (Figure 3B1), whereas when a 5 &#181;L/s flowrate is applied, the spheres move through the field of view, as evidenced by streaking (Figure 3B2). This same approach was used to induce flow through a 2D, vascular-derived network to monitor flow of TRITC-labeled, 65 kDa dextran through the network and diffusion into the surrounding hydrogel (Figure 3C).
<img alt="Figure 3" src="/files/ftp_upload/55101/55101fig3.jpg" /> 
Figure 3: Syringe Pump-Driven Flow in PEGDA Hydrogels Polymerized in Microfluidic Housings. A commercially available, pre-fabricated microfluidic device (A) with a 17 x 3.8 x 0.53 mm (x,y,z) microchannel houses a photopolymerized 1 x 3.8 x 0.53 mm (x,y,z) PEGDA hydrogel, indicated by the black arrow. A 20-&#181;m diameter cylindrical channel was degraded through the hydrogel, and 2-&#181;m polystyrene spheres were pumped through the device (B). The spheres are clearly resolved without fluid flow (B1), but they appear as streaks at a flowrate of 5 &#181;L/s (B2). In another housed hydrogel, a 2D vascular-derived network was fabricated and TRITC-labeled, 65 kDa dextran was pumped through the network (C). Time-lapse microscopy was used to monitor the fluorescent dextran as it filled the channels and diffused into the surrounding hydrogel. White arrows in (B) and (C) indicate the direction of fluid flow. The calibration bar in (C) indicates the intensity of the fluorescent dextran. The scale bars represent 20 &#181;m in (B1) and (B2) and 50 &#181;m in (C). <a href="https://cloudflare-jove-com.bdigitaluss.remotexs.co/files/ftp_upload/55101/55101fig3large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>

Watch this Scientific Journal Video about Image-guided, Laser-based Fabrication of Vascular-derived Microfluidic Networks at JoVE.com

Image-guided, Laser-based Fabrication of Vascular-derived Microfluidic Networks

This protocol outlines the implementation of image-guided, laser-based hydrogel degradation to fabricate vascular-derived, biomimetic microfluidic networks embedded in poly(ethylene glycol) diacrylate (PEGDA) hydrogels. These biomimetic microfluidic systems may be useful for tissue engineering applications, generation of in vitro disease models, and fabrication of advanced "on-a-chip" devices.

This detailed protocol outlines the implementation of image-guided, laser-based hydrogel degradation for the fabrication of vascular-derived microfluidic ...

image-guided-laser-based-fabrication-vascular-derived-microfluidic

Universidad San Sebastian

Research

JoVE Journal

Bioengineering

9.8K Views.  University of Delaware. This protocol outlines the implementation of image-guided, laser-based hydrogel degradation to fabricate vascular-derived, biomimetic microfluidic networks embedded in poly(ethylene glycol) diacrylate (PEGDA) hydrogels. These biomimetic microfluidic systems may be useful for tissue engineering applications, generation of in vitro disease models, and fabrication of advanced "on-a-chip" devices.

Video: Image-guided, Laser-based Fabrication of Vascular-derived Microfluidic Networks

Procedure for the Development of Multi-depth Circular Cross-sectional Endothelialized Microchannels-on-a-chip

Efforts have been focused on developing in vitro assays for the study of microvessels because in vivo animal studies are more time-consuming, expensive, and observation and quantification are very challenging. However, conventional in vitro microvessel assays have limitations when representing in vivo microvessels with respect to three-dimensional (3D) geometry and providing continuous fluid flow. Using a combination of photolithographic reflowable photoresist technique, soft lithography, and microfluidics, we have developed a multi-depth circular cross-sectional endothelialized microchannels-on-a-chip, which mimics the 3D geometry of in vivo microvessels and runs under controlled continuous perfusion flow. A positive reflowable photoresist was used to fabricate a master mold with a semicircular cross-sectional microchannel network. By the alignment and bonding of the two polydimethylsiloxane (PDMS) microchannels replicated from the master mold, a cylindrical microchannel network was created. The diameters of the microchannels can be well controlled. In addition, primary human umbilical vein endothelial cells (HUVECs) seeded inside the chip showed that the cells lined the inner surface of the microchannels under controlled perfusion lasting for a time period between 4 days to 2 weeks.

A microchannels-on-a-chip platform was developed by the combination of photolithographic reflowable photoresist technique, soft lithography, and microfluidics. The endothelialized microchannels platform mimics the three-dimensional (3D) geometry of in vivo microvessels, runs under controlled continuous perfusion flow, allows for high-quality and real-time imaging&#160;and can be applied for microvascular research.

Efforts have been focused on developing in vitro assays for the study of microvessels because in vivo animal studies are more time-consuming, expensive, ...

Microfluidic Genipin Deposition Technique for Extended Culture of Micropatterned Vascular Muscular Thin Films

The chronic nature of vascular disease progression requires the development of experimental techniques that simulate physiologic and pathologic vascular behaviors on disease-relevant time scales. Previously, microcontact printing has been used to fabricate two-dimensional functional arterial mimics through patterning of extracellular matrix protein as guidance cues for tissue organization. Vascular muscular thin films utilized these mimics to assess functional contractility. However, the microcontact printing fabrication technique used typically incorporates hydrophobic PDMS substrates. As the tissue turns over the underlying extracellular matrix, new proteins must undergo a conformational change or denaturing in order to expose hydrophobic amino acid residues to the hydrophobic PDMS surfaces for attachment, resulting in altered matrix protein bioactivity, delamination, and death of the tissues.
Here, we present a microfluidic deposition technique for patterning of the crosslinker compound genipin. Genipin serves as an intermediary between patterned tissues and PDMS substrates, allowing cells to deposit newly-synthesized extracellular matrix protein onto a more hydrophilic surface and remain attached to the PDMS substrates. We also show that extracellular matrix proteins can be patterned directly onto deposited genipin, allowing dictation of engineered tissue structure. Tissues fabricated with this technique show high fidelity in both structural alignment and contractile function of vascular smooth muscle tissue in a vascular muscular thin film model. This technique can be extended using other cell types and provides the framework for future study of chronic tissue- and organ-level functionality.

We present a method for microfluidic deposition of patterned genipin and fibronectin on PDMS substrates, allowing extended viability of vascular smooth muscle cell-dense tissues. This tissue fabrication method is combined with previous vascular muscular thin film technology to measure vascular contractility over disease-relevant time courses.

The chronic nature of vascular disease progression requires the development of experimental techniques that simulate physiologic and pathologic vascular ...

Micropatterning and Assembly of 3D Microvessels

In vitro platforms to study endothelial cells and vascular biology are largely limited to 2D endothelial cell culture, flow chambers with polymer or glass based substrates, and hydrogel-based tube formation assays. These assays, while informative, do not recapitulate lumen geometry, proper extracellular matrix, and multi-cellular proximity, which play key roles in modulating vascular function. This manuscript describes an injection molding method to generate engineered vessels with diameters on the order of 100 &#181;m. Microvessels are fabricated by seeding endothelial cells in a microfluidic channel embedded within a native type I collagen hydrogel. By incorporating parenchymal cells within the collagen matrix prior to channel formation, specific tissue microenvironments can be modeled and studied. Additional modulations of hydrodynamic properties and media composition allow for control of complex vascular function within the desired microenvironment. This platform allows for the study of perivascular cell recruitment, blood-endothelium interactions, flow response, and tissue-microvascular interactions. Engineered microvessels offer the ability to isolate the influence from individual components of a vascular niche and precisely control its chemical, mechanical, and biological properties to study vascular biology in both health and disease.

This manuscript presents an injection molding method to engineer microvessels that recapitulate physiological properties of endothelium. The microfluidic-based process creates patent 3D vascular networks with tailorable conditions, such as flow, cellular composition, geometry, and biochemical gradients. The fabrication process and examples of potential applications are described.

In vitro platforms to study endothelial cells and vascular biology are largely limited to 2D endothelial cell culture, flow chambers with polymer or glass ...

Preparing a Microfluidics-Based Vascularized Human Micro-Tumor Model

Establishing a Physiologic Human Vascularized Micro-Tumor Model for Cancer Research

A lack of validated cancer models that recapitulate the tumor microenvironment of solid cancers in vitro remains a significant bottleneck for preclinical cancer research and therapeutic development.&#160;To overcome this problem, we have developed the vascularized microtumor (VMT), or tumor chip, a microphysiological system that realistically models the complex human tumor microenvironment. The VMT forms de novo within a microfluidic platform by co-culture of multiple human cell types under dynamic, physiological flow conditions. This tissue-engineered micro-tumor construct incorporates a living perfused vascular network that supports the growing tumor mass just as newly formed vessels do in vivo. Importantly, drugs and immune cells must cross the endothelial layer to reach the tumor, modeling in vivo physiological barriers to therapeutic delivery and efficacy. Since the VMT platform is optically transparent, high-resolution imaging of dynamic processes such as immune cell extravasation and metastasis can be achieved with direct visualization of fluorescently labeled cells within the tissue. Further, the VMT retains in vivo tumor heterogeneity, gene expression signatures, and drug responses. Virtually any tumor type can be adapted to the platform, and primary cells from fresh surgical tissues grow and respond to drug treatment in the VMT, paving the way toward truly personalized medicine. Here, the methods for establishing the VMT and utilizing it for oncology research are outlined. This innovative approach opens new possibilities for studying tumors and drug responses, providing researchers with a powerful tool to advance cancer research.

Author Spotlight: Creating Human Vascularized Micro-Tumors as Models for Translational Cancer Research 

This protocol presents a physiologically relevant tumor-on-a-chip model to perform high-throughput basic and translational human cancer research, advancing drug screening, disease modeling, and personalized medicine approaches with a description of loading, maintenance, and evaluation procedures.

A lack of validated cancer models that recapitulate the tumor microenvironment of solid cancers in vitro remains a significant bottleneck for preclinical ...

Cancer Research

Microfluidic Bioprinting for Engineering Vascularized Tissues and Organoids

Engineering vascularized tissue constructs and organoids has been historically challenging. Here we describe a novel method based on microfluidic bioprinting to generate a scaffold with multilayer interlacing hydrogel microfibers. To achieve smooth bioprinting, a core-sheath microfluidic printhead containing a composite bioink formulation extruded from the core flow and the crosslinking solution carried by the sheath flow, was designed and fitted onto the bioprinter. By blending gelatin methacryloyl (GelMA) with alginate, a polysaccharide that undergoes instantaneous ionic crosslinking in the presence of select divalent ions, followed by a secondary photocrosslinking of the GelMA component to achieve permanent stabilization, a microfibrous scaffold could be obtained using this bioprinting strategy. Importantly, the endothelial cells encapsulated inside the bioprinted microfibers can form the lumen-like structures resembling the vasculature over the course of culture for 16 days. The endothelialized microfibrous scaffold may be further used as a vascular bed to construct a vascularized tissue through subsequent seeding of the secondary cell type into the interstitial space of the microfibers. Microfluidic bioprinting provides a generalized strategy in convenient engineering of vascularized tissues at high fidelity.

We provide a generalized protocol based on a microfluidic bioprinting strategy for engineering a microfibrous vascular bed, where a secondary cell type could be further seeded into the interstitial space of this microfibrous structure to generate vascularized tissues and organoids.

Engineering vascularized tissue constructs and organoids has been historically challenging. Here we describe a novel method based on microfluidic ...

Perfusable Vascular Network with a Tissue Model in a Microfluidic Device

A spheroid (a multicellular aggregate) is regarded as a good model of living tissues in the human body. Despite the significant advancement in the spheroid cultures, a perfusable vascular network in the spheroids remains a critical challenge for long-term culture required to maintain and develop their functions, such as protein expressions and morphogenesis. The protocol presents a novel method to integrate a perfusable vascular network within the spheroid in a microfluidic device. To induce a perfusable vascular network in the spheroid, angiogenic sprouts connected to microchannels were guided to the spheroid by utilizing angiogenic factors from human lung fibroblasts cultured in the spheroid. The angiogenic sprouts reached the spheroid, merged with the endothelial cells co-cultured in the spheroid, and formed a continuous vascular network. The vascular network could perfuse the interior of the spheroid without any leakage. The constructed vascular network may be further used as a route for supply of nutrients and removal of waste products, mimicking blood circulation in vivo. The method provides a new platform in spheroid culture toward better recapitulation of living tissues.

The protocol describes how to engineer a perfusable vascular network in a spheroid. The spheroid's surrounding microenvironment is devised to induce angiogenesis and connect the spheroid to the microchannels in a microfluidic device. The method allows the perfusion of the spheroid, which is a long-awaited technique in three-dimensional cultures.

A spheroid (a multicellular aggregate) is regarded as a good model of living tissues in the human body. Despite the significant advancement in the ...

Stepwise Cell Seeding on Tessellated Scaffolds to Study Sprouting Blood Vessels

The cardiovascular system is a key player in human physiology, providing nourishment to most tissues in the body; vessels are present in different sizes, structures, phenotypes, and performance depending on each specific perfused tissue. The field of tissue engineering, which aims to repair or replace damaged or missing body tissues, relies on controlled angiogenesis to create a proper vascularization within the engineered tissues. Without a vascular system, thick engineered constructs cannot be sufficiently nourished, which may result in cell death, poor engraftment, and ultimately failure. Thus, understanding and controlling the behavior of engineered blood vessels is an outstanding challenge in the field. This work presents a high-throughput system that allows for the creation of organized and repeatable vessel networks for studying vessel behavior in a 3D scaffold environment. This two-step seeding protocol shows that vessels within the system react to the scaffold topography, presenting distinctive sprouting behaviors depending on the compartment geometry in which the vessels reside. The obtained results and understanding from this high throughput system can be applied in order to inform better 3D bioprinted scaffold construct designs, wherein fabrication of various 3D geometries cannot be rapidly assessed when using 3D printing as the basis for cellularized biological environments. Furthermore, the understanding from this high throughput system may be utilized for the improvement of rapid drug screening, the rapid development of co-cultures models, and the investigation of mechanical stimuli on blood vessel formation to deepen the knowledge of the vascular system.

Engineered tissues heavily rely on proper vascular networks to provide vital nutrients and gases and remove metabolic waste. In this work, a stepwise seeding protocol of endothelial cells and support cells creates highly organized vascular networks in a high-throughput platform for studying developing vessel behavior in a controlled 3D environment.

The cardiovascular system is a key player in human physiology, providing nourishment to most tissues in the body; vessels are present in different sizes, ...

A Microphysiological System to Study Leukocyte-Endothelial Cell Interaction during Inflammation

Leukocyte-endothelial cell interactions play an important role in inflammatory diseases such as sepsis. During inflammation, excessive migration of activated leukocytes across the vascular endothelium into key organs can lead to organ failure. A physiologically relevant biomimetic microfluidic assay (bMFA) has been developed and validated using several experimental and computational techniques, which can reproduce the entire leukocyte rolling/adhesion/migration cascade to study leukocyte-endothelial cell interactions. Microvascular networks obtained from in vivo images in rodents were digitized using a Geographic Information System (GIS) approach and microfabricated with polydimethylsiloxane (PDMS) on a microscope slide. To study the effect of shear rate and vascular topology on leukocyte-endothelial cell interactions, a Computational Fluid Dynamics (CFD) model was developed to generate a corresponding map of shear rates and velocities throughout the network. The bMFA enables the quantification of leukocyte-endothelial cells interactions, including rolling velocity, number of adhered leukocytes in response to different shear rates, number of migrated leukocytes, endothelial cell permeability, adhesion molecule expression and other important variables. Furthermore, by using human-related samples, such as human endothelial cells and leukocytes, bMFA provides a tool for rapid screening of potential therapeutics to increase their clinical translatability.

In this protocol, a biomimetic microfluid assay, which can reproduce a physiologically relevant microvascular environment and reproduce the entire leukocyte adhesion/migration cascade, is employed to study leukocyte-endothelial cell interactions in inflammatory disease.

Leukocyte-endothelial cell interactions play an important role in inflammatory diseases such as sepsis. During inflammation, excessive migration of ...

Fabrication of Engineered Vascular Flaps Using 3D Printing Technologies

Engineering implantable, functional, thick tissues requires designing a hierarchical vascular network. 3D bioprinting is a technology used to create tissues by adding layer upon layer of printable biomaterials, termed bioinks, and cells in an orderly and automatic manner, which allows for creating highly intricate structures that traditional tissue engineering techniques cannot achieve. Thus, 3D bioprinting is an appealing in vitro approach to mimic the native vasculature complex structure, ranging from millimetric vessels to microvascular networks. 
Advances in 3D bioprinting in granular hydrogels enabled the high-resolution extrusion of low-viscosity extracellular matrix-based bioinks. This work presents a combined 3D bioprinting and sacrificial mold-based 3D printing approach for fabricating engineered vascularized tissue flaps. 3D bioprinting of endothelial and support cells using recombinant collagen-methacrylate bioink within a gelatin support bath is utilized for the fabrication of a self-assembled capillary network. This printed microvasculature is assembled around a mesoscale vessel-like porous scaffold, fabricated using a sacrificial 3D printed mold, and is seeded with endothelial cells. 
This assembly induces the endothelium of the mesoscale vessel to anastomose with the surrounding capillary network, establishing a hierarchical vascular network within an engineered tissue flap. The engineered flap is then directly implanted by surgical anastomosis to a rat femoral artery using a cuff technique. The described methods can be expanded for the fabrication of various vascularized tissue flaps for use in reconstruction surgery and vascularization studies.

Engineered flaps require an incorporated functional vascular network. In this protocol, we present a method of fabricating a 3D printed tissue flap containing a hierarchical vascular network and its direct microsurgical anastomoses to rat femoral artery.

Engineering implantable, functional, thick tissues requires designing a hierarchical vascular network. 3D bioprinting is a technology used to create ...

3D Bioprinting for Rapid Vascular Network Fabrication Using Vessels-on-a-Plate

High-Throughput Bioprinting Method for Modeling Vascular Permeability in Standard Six-well Plates with Size and Pattern Flexibility

Vascular permeability is a key factor in developing therapies for disorders associated with compromised endothelium, such as endothelial dysfunction in coronary arteries and impaired function of the blood-brain barrier. Existing fabrication techniques do not adequately replicate the geometrical variation in vascular networks in the human body, which substantially influences disease progression; moreover, these techniques often involve multi-step fabrication procedures that hinder the high-throughput production necessary for pharmacological testing. This paper presents a bioprinting protocol for creating multiple vascular tissues with desired patterns and sizes directly on standard six-well plates, overcoming existing resolution and productivity challenges in bioprinting technology. A simplified fabrication approach was established to construct six hollow, perfusable channels within a hydrogel, which were subsequently lined with human umbilical vein endothelial cells to form a functional and mature endothelium. The computer-controlled nature of 3D bioprinting ensures high reproducibility and requires fewer manual fabrication steps than traditional methods. This highlights VOP's potential as an efficient high-throughput platform for modeling vascular permeability and advancing drug discovery.

Author Spotlight: Automated Bioprinting for High-Throughput Vascular Model Fabrication

We present a protocol for high-throughput production of vascular channels with flexible sizes and desired patterns on a standard six-well plate using 3D bioprinting technology, referred to as vessels-on-a-plate (VOP). This platform has the potential to advance the development of therapeutics for the disorders associated with compromised endothelium.

Vascular permeability is a key factor in developing therapies for disorders associated with compromised endothelium, such as endothelial dysfunction in ...