This work was supported by National Institutes of Health National Cancer Institute SBIR Phase I (HHSN26120700015C) and P01CA098912.

Tumor tissue was obtained with patient consent and according to an approved Institutional Review Board (IRB) protocol. Xenografts were implanted, grown, and harvested according to an accepted Institutional Animal Care and Use Committee (IACUC) protocol.
NOTE: All work is to be performed in a sterile biological safety cabinet to maintain sterility. All steps should be conducted at room temperature unless otherwise specified.
1. Preparation of materials for PDX processing
<ol>
	<li>Autoclave forceps and scalpel handle or razor blades.</li>
	<li>Thaw dissociation enzyme solution at 4 &#176;C overnight or at room temperature the same day as tissue dissociation. 
	NOTE: Thawing at 37 &#176;C is not advised, as this can inactivate some dissociation enzymes.</li>
	<li>Prepare at least 100 mL of PDX culture medium (Dulbecco&#8217;s modified Eagle medium-nutrient mixture F-12 [DMEM-F12] with 100 U/mL penicillin and streptomycin and 30% fetal bovine serum [FBS]), and at least 25 mL of PDX processing medium (DMEM-F12 with 100&#160;U/mL penicillin and streptomycin and no FBS). Store at 4 &#176;C until ready to use.</li>
</ol>
2. PDX dissociation and initial purification of stromal component
<ol>
	<li>Gather autoclaved utensils, 70 &#181;m cell strainers (2&#8722;3), 60 mm round tissue culture dishes (2), 6-well tissue culture plates, sterile 50 mL conical centrifuge tubes, and sterile 1x phosphate-buffered saline (PBS). Warm culture medium to 37 &#176;C and allow dissociation enzyme solution to come to room temperature.</li>
	<li>When PDX tissue has reached a diameter of 1.0&#8722;1.5 cm in a mouse host, surgically remove the tumor from the mouse by standard means (e.g., under accepted anesthesia) and store on ice in a 50 mL tube with PDX culture medium (Figure 2A). Process the tissue promptly, via the steps below, to maximize cell viability, preferably within 1&#8722;2 h after harvest.</li>
	<li>Transfer tumor tissue to a pre-weighed sterile 50 mL conical tube. Rinse 6x with 30&#160;mL of sterile PBS to remove blood and contaminants. Remove as much liquid as possible and weigh the tumor tissue.</li>
	<li>Transfer tumor tissue to a 60 mm round tissue culture dish and mince into ~1 mm3 pieces using a sterile razor blade or scalpel.</li>
	<li>Add 5 mL of PDX processing medium to collect tumor slurry and transfer to a new sterile 50 mL tube. Rinse the culture dish with another 5 mL of PDX processing medium, then with dissociation enzyme solution (10 mL/g tumor, at least 5 mL), adding all rinses to the 50 mL tube.</li>
	<li>Incubate 20 min at 37 &#176;C with gentle shaking. Swirl the tube gently halfway through the incubation time.</li>
	<li>Pipette up and down gently with a serological pipette to break up clumps. Filter cells with a 70 &#181;m cell strainer placed over a new sterile 50 mL tube. 
	NOTE: More than one strainer may be necessary.</li>
	<li>Centrifuge at 200 x g for 5 min to pellet cells. Remove supernatant and resuspend in 2&#8722;3 mL of PDX culture medium. Count cells using a hemocytometer or automated cell counter.</li>
	<li>Use Table 1 to estimate the required number of dissociated PDX-derived cells needed to achieve the desired cell density per chip. 
	NOTE: The values in Table 1 are intended to be a starting point. Actual values will vary due to tissue viability/cellularity and cell loss from freezing/recovery. PDX tumors are individuals even within a given cancer type so these values should be adjusted empirically.</li>
	<li>Of the number calculated in step 2.9, plate 1&#8722;2 x 106 cells in 5 mL of PDX culture medium per well of a 6-well tissue culture plate. Incubate (37&#160;&#176;C, 5% CO2, 95% humidity) for 48 h with gentle shaking (50&#8722;55 rpm) to promote cluster formation (Figure 2B). After cluster have formed, proceed to section 3 for centrifugation.</li>
	<li>Cryopreserve unused PDX cells from the initial dissociation in 50% FBS + 40% DMEM-F12 + 10% dimethyl sulfoxide (DMSO) or a commercially available primary cell freezing medium. 
	NOTE: Adherent mouse stromal cells may also be recovered from the tissue plate surface, if desired, by rinsing briefly with culture medium and expanding by standard means.</li>
	<li>For later use of cryopreserved PDX tumor cells/mouse stroma, thaw cells in a 37 &#176;C water bath for 2 min. Count and plate in a 6-well tissue culture plate as described in step 2.10 before proceeding to section 3. Increase the number of PDX cells by ~20% to accommodate viability losses from cryopreservation.</li>
</ol>
3. Density gradient centrifugation-based separation of PDX-derived clusters from single cells
<ol>
	<li>Prepare 20 mL of 100% density gradient solution by thoroughly mixing 18 mL of density gradient centrifugation solution with 2 mL of sterile 10x Hanks&#8217; balanced salt solution (HBSS) in a sterile 50 mL conical tube. Make 10 mL each of 20%, 30%, 40%, and 55% density gradient solutions by diluting this 100% solution with sterile 1x HBSS and mixing well. 
	NOTE: These volumes are sufficient for two 15 mL gradients which can be used to separate ~15 x 106 cells each. If separating fewer than ~15 x 106 cells, the second gradient should be used as a balance for centrifugation.</li>
	<li>Add 3 mL of 55% density gradient solution to the bottom of a 15 mL conical tube. Holding the tube at an angle, very gently layer 3 mL of 40% density gradient solution on top of the 55% layer, slowly dispensing the liquid onto the angled side of the tube to avoid mixing the layers. Repeat with the 30% density gradient solution.</li>
	<li>Collect the supernatant of PDX rotation cultures with a 5 mL serological pipette, rinsing plate surface gently. Centrifuge at 200 x g for 2 min to pellet cells.</li>
	<li>Remove supernatant and resuspend the cell pellet in 3 mL of 20% density gradient solution according to number of gradients needed to separate the cells. Carefully layer 20% density gradient solution with cells onto the top of the gradient(s). If only using one gradient tube for cells, top the balance gradient tube with cell-free 20% density gradient solution.</li>
	<li>Cap the tubes and centrifuge in a swing bucket rotor centrifuge for 30 min at 4 &#176;C, 2,000 x g, and 0 brake.</li>
	<li>After centrifugation, fractions will be visible (Figure 2C). Collect 2&#8722;3 mL of fractions into fresh 15 mL tubes. Add 3&#8722;4 volumes of sterile 1x HBSS to each fraction and invert to mix thoroughly.</li>
	<li>Centrifuge at 1,000 x g for 3 min to pellet cells. Remove supernatant and resuspend the cell pellet in 1&#8722;2 mL of PDX processing medium. 
	NOTE: Viable PDX cell clusters are typically found at the 40&#8722;55% density gradient solution interface (Figure 2D) with single dead/dying cells accumulating at the 20&#8722;40% density gradient solution interface for most PDXs tested.</li>
	<li>Remove a small, representative aliquot (50&#8722;100 &#181;L) for re-dissociation with an equal volume of dissociation enzyme solution to assess cell number in the clustered cell suspension. Count cells with a hemocytometer or automated cell counter.</li>
</ol>
4. Hydrogel preparation and microfluidic plate seeding
<ol>
	<li>Reconstitute HA hydrogel solutions (thiol-modified HA, HA-SH; thiol-reactive polyethylene glycol diacrylate, PEGDA) according to the manufacturer&#8217;s instructions.</li>
	<li>Using a multichannel pipette, add 50 &#956;L of HBSS to all wells in observation window columns (column 3, 7, 11, 15, 19, 23) of a 2-lane microfluidic plate to maintain culture humidity and optimal imaging conditions.</li>
	<li>Calculate the volume of cell suspension from section 3 needed for 50 &#956;L of hydrogel at the desired cell density (i.e., 5,000 cells/&#956;L). For seeding one microfluidic plate, aliquot the calculated volume into each of 4 sterile 1.5 mL centrifuge tubes. 
	NOTE: HA hydrogels have a fixed time to gelation. Adjust the volume of gel solution aliquots for user efficiency at dispensing if premature gelation occurs.</li>
	<li>Adjust the pH of the HA-SH solution to 8.0 with 1 N NaOH immediately prior to use. Perform a test gelation by mixing 40 &#956;L of HA-SH with 10 &#956;L of PEGDA and monitoring gelation over time. Gelation typically begins 5&#8722;8 min after mixing HA-SH with the PEGDA crosslinker.</li>
	<li>Centrifuge cell suspension aliquots for 2 min (200 x g, room temperature) to pellet cells. Carefully remove the supernatant and resuspend cells in the appropriate volume of HA-SH. 
	NOTE: Final hydrogel is a 4:1 HA-SH:PEGDA solution (by volume) so cells should be resuspended in 40 &#956;L of HA-SH for a 50 &#956;L final volume.</li>
	<li>Add 10 &#956;L of PEGDA to one aliquot of cells in HA. Mix well and wait 1&#8722;3 min (depending on gelation time from step 4.4) before seeding the microfluidic plate. 
	NOTE: Allowing the gelation reaction of start before seeding helps to minimize cell settling.</li>
	<li>Affix a tip for dispensing 1.5 &#956;L volumes to a single channel repeating pipette and load with cells in HA hydrogel solution. Remember to keep the hydrogel aliquot well-mixed to ensure even cell distribution.</li>
	<li>To seed the microfluidic plate, align pipette tip perpendicular to the plate while gently placing the tip in the center of the gel inlet (columns 1, 5, 9, 13, 17, 21) to ensure contact but no pressure when dispensing the hydrogel solution. Working quickly to prevent premature hydrogel solidification, dispense 1.5 &#956;L of gel solution into each gel inlet.</li>
	<li>Observe the fill status of the microfluidic channels by viewing from the top of the plate, bottom of the plate, or by microscope, and assess loading, using Figure 3 as a guide (successful loading in Figure 3A, pipet positioning guidance in Figure 3B, missed loading in Figure 3C, not filled to end in Figure 3D, overflow in Figure 3E). Identify any necessary adjustments in technique that may improve filling success for the next round of chips (see discussion for troubleshooting tips).</li>
	<li>Repeat steps 4.6&#8722;4.9 with the remaining 3 aliquots of cells in HA solution. Invert the plate while preparing the next aliquot (~1 min). 
	NOTE: The 1 min wait time and inversion of the plate improve the 3D distribution of the cells by reducing cell settling as gelation occurs.</li>
	<li>After all chips are filled, incubate the plate at 37 &#176;C in a humidified incubator until gelation is complete (~45 min).</li>
	<li>Using the manual provided, ensure the perfusion rocker is installed in the cell culture incubator with the correct perfusion settings (14&#176; angle, 4 min intervals).</li>
	<li>Add 50 &#956;L of PDX culture medium to all medium inlets (columns 2, 6, 10, 14, 18, 22) and check if the channels filled properly by flipping the plate. Gently tap the plate against a surface to encourage the liquid to fill the microfluidic channels.</li>
	<li>Add 50 &#956;L of DMEM-F12 (10% FBS) for all medium outlets (column 4, 8, 12, 16, 20, 24). If any air bubbles are trapped in the perfusion channel, remove by gently tapping the plate against a surface.</li>
	<li>Using a microscope and plate layout form (Supplemental Figure 1), record chip filling success. Exclude improperly filled chips from further experimental use.</li>
	<li>Place plate on a tilting rocker set to a 14&#176; tilt and a 4 min cycle to begin perfusion. Replace PDX culture medium every 2 days (first 50 &#956;L in inlet, then 50 &#956;L in outlet).</li>
</ol>
5. Cell staining, imaging, and image quantification
<ol>
	<li>Prepare a cell viability assay solution containing three fluorescent dyes (Hoechst 33342, ethidium homodimer-1, calcein acetoxymethyl [AM]).
	<ol>
		<li>Prepare stock solutions of each dye as follows: Hoechst 33342 at 1.6 mM (1 mg/mL) in deionized water; calcein AM at 4 mM in anhydrous DMSO; ethidium homodimer-1 at 2 mM in DMSO/H2O (1:4, v/v). 
		NOTE: The stock calcein AM and ethidium homodimer-1 solutions are provided in the noted kit in Table of Materials.</li>
		<li>Prepare a single working solution in HBSS or phenol red-free medium, containing all three dyes. Optimize final working concentration for each cell type and matrix, within the ranges of 1.6&#8722;8.0 &#181;M for Hoechst 33342, 0.1&#8722;10 &#181;M for calcein AM, and 0.1&#8722;10 &#181;M for ethidium homodimer-1.</li>
	</ol>
	</li>
	<li>Remove culture medium and apply the working viability dye solution to desired microfluidic chips (75 &#956;L in inlet, 25 &#956;L in outlet) and place back on the perfusion rocker in the cell culture incubator for 1 h.</li>
	<li>Image the observation windows of the stained cultures using a manual or automated confocal microscope (Table of Materials) with fluorescent filters (listed as excitation/emission wavelengths, in nm) to observe all nuclei (Hoechst 33342, 350/461), dead cell nuclei (ethidium homodimer-1, 528/617), and live cell cytoplasm (calcein AM, 494/517).</li>
	<li>Capture 140 &#181;m Z-stacks with a step size of &#8804;1 &#181;m using a 20x air objective. Three fields of view are needed to image the entire microchannel with a small amount of overlap. To avoid double sampling, image only two fields of view per chip. 
	NOTE: Imaging conditions should be optimized to ensure proper NyQuist sampling. In authors&#8217; experience, a fast imaging system, based either on deconvolution of a conventional epi-fluorescence source or resonance scanning mode on a confocal, is necessary to fully assay a complete Z-stack, with three laser colors, across 96 chips on a plate within a reasonable amount of time (roughly 3.5 h with automated imaging, including setup).</li>
	<li>Using image analysis software, assay the Z-stack images for the desired quantified data, such as morphology, aggregation state, or other features. To quantify cell viability, count the number of dead cells (red) and total cell nuclei (blue).</li>
</ol>

<ol>
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The authors have nothing to disclose.

Here we describe a method for processing and culturing viable PDX-derived tumor cells in a high-throughput, perfused 3D culture system. While this protocol utilizes PCa PDX tissue, it is equally effective for other epithelial-derived tumors. Tumor characteristics vary between individual PDX lines even within the same tissue of origin (prostate, breast, etc.). Some PCa PDX lines are more fibrotic and difficult to isolate viable cells from while others are more cellular. The tumor size noted here can be varied within IACUC...

Over the past decade, the field of cancer research has demonstrated renewed enthusiasm for patient-derived xenografts (PDXs) as a tool for assessing cancer cell pathway reliance and drug susceptibility1. The most common PDX models are established by subcutaneous or orthotopic implantation of human tumor cells&#8212;a tumor fragment, a cluster of dissociated tumor-derived cells, or a sample of isolated circulating tumor cells (CTCs)&#8212;into a rodent host. If the tumor &#8220;take&#8221; is successful, the xenograft cells will proliferate, vascularize, and otherwise interact with the host tissue to create a tumor, which can be harvested at an optimal size, subdivided, and re-implanted into other hosts. Among their many advantages as a model system, PDXs typically retain a substantial portion of the native tumor cell population&#8217;s heterogeneity and enable the assessment of human-specific pathways and cell responses2,3. The in vivo context enables tumor interaction with vasculature and other adjacent stroma and recapitulates tissue characteristics such as drug diffusion dynamics, oxygen tension, and extracellular matrix influence that biologically and mechanically impact tumor progression. A negative aspect of PDXs is their reliance on a rodent host, both for tumor expansion and ultimately for hypothesis testing. Because many PDXs cannot adapt to traditional two-dimensional (2D) culture on tissue culture polystyrene without losing many of their desirable characteristics, there has been minimal middle ground for researchers between this relatively controlled in vitro method, and the significant increase in expense, facilities, and time requirements for in vivo PDX use.
We have described multiple in vitro models that implement 3D cell culture within a supportive matrix, and recently expanded that work to demonstrate the ex vivo culture of multiple prostate cancer (PCa)-derived PDXs, both alone and in co-culture with bone marrow-derived fibroblasts4,5. Hyaluronic acid (HA)-based hydrogel matrices provide customizable and biologically-relevant support for both cell types, with facile control over hydrogel characteristics and optical clarity for imaging through the hydrogel depth6.
Mature PDX tumor tissues comprise a variable mixture of heterogeneous human cancer cells and mouse stroma (fibroblasts, endothelial cells, etc.). To study cell-type specific contributions to tumor progression in vitro, it can be advantageous to dissociate tumors, separate the cell populations, and experimentally incorporate them in an organized manner to dissect pathways of intercellular communication. The mixed cell populations within tissue digestates have differential compatibility with specific culture conditions. For example, tumor-associated fibroblast viability necessitates either surface adherence or 3D matrices functionalized with integrin ligands, while epithelial-derived PDX cells do not typically have these requirements, instead favoring cell-cell interactions. These differences can be exploited to achieve effective separation of PDX cells from contaminating mouse stromal cells. Rotation culture of tissue digestates allows stromal cell adherence to the tissue culture surface while cell-cell adhesions drive PDX cells floating above the rotating culture surface to form multicellular clusters in the supernatant in 24&#8722;48 h. The specific characteristics of these clusters vary with the PDX (e.g., large, tight, highly spherical clusters or smaller, looser aggregates resembling bunches of grapes), but are typically of biologically relevant sizes (50&#8722;250 &#181;m diameter), sufficient for assessing cellular interactions that rely on intercellular contacts.
Tumor retrieval and processing inevitably results in some degree of collateral cell death, either due to short-term damage from mechanical/enzymatic disruption, or long-term incompatibility of subpopulations with the chosen culture conditions. Despite the utility of rotation culture as an initial bulk separation, dead or dying cells are inevitably transferred with the PDX clusters and can influence the resultant culture. These dead cells are often individual PDX cells that were not integrated into a cluster, mouse stromal fibroblasts that cannot survive in selected culture conditions, or particularly fragile endothelial cells. Such dying cells can influence experimental results from &#8220;survivors&#8221; and can substantially impact quantification, e.g., via fluorescent image-based viability screening assays. To improve the selection of live PDX cells from this method, we adapted centrifugation methods with density steps to easily remove individual dead/dying cells from PDX mixtures and retain predominantly live multicellular clusters.
To enhance the study of resultant PDX-derived clusters in 3D culture, we utilized a microfluidics-based perfusion culture platform, the OrganoPlate (Figure 1), which is a high-throughput organ-on-a-chip platform that allows for simultaneous culture of up to 96 individual perfused, 3D cultures on a 384-well microtiter plate-base (Figure 1A)7,8. In the 2-lane microfluidic plate, a single tissue chip is connected by two microfluidic channels (Figure 1B, gel channel: red, perfusion channel: blue) which span four wells in a row. The two microfluidic channels are separated by a short plastic ridge called a Phaseguide which prevents overflow of one channel into its adjacent neighbor channel, and simultaneously allows for a membrane-free interface between the contents of the gel and perfusion channel9. Because the bottom of the microfluidic plate is composed of microscope-grade glass, the cultures can be viewed in the observation window through the bottom of the plate with a standard or automated microscope. Perfusion is established in the microfluidic plate with a programmable rocker, using gravity to drive media through the microfluidic channels, between reservoir wells (Figure 1C). The perfusion flow-mimic more closely recapitulates the tumor microenvironment than static culture, allowing for the incorporation of shear stress and enhanced distribution of gases and nutrients. The benefits of maintaining a perfused cancer cell culture in the microfluidic plate have previously been described as perfused breast cancer cultures exhibited optimal viability as compared to a static 3D culture of the same cells7.
The present report describes an adapted density gradient centrifugation method for isolating live multicellular PDX clusters and demonstrates its utility in establishing 3D PDX cultures within perfusable microfluidic plates. Because an increasing number of research laboratories are seeking methods to facilitate PDX use, we anticipate that the protocols presented here will be of immediate utility.

<table><tbody><tr><td>1N NaOH</td><td></td><td></td><td>any suitable tissue culture grade</td></tr><tr><td>60 mm round tissue culture dishes</td><td></td><td></td><td>any suitable</td></tr><tr><td>6-well tissue culture plates</td><td></td><td></td><td>any suitable</td></tr><tr><td>70 &amp;micro;m cell strainers</td><td>Corning</td><td>431751</td><td>or equivalent</td></tr><tr><td>Centrifuge</td><td>Eppendorf</td><td>5810R with suitable rotor and buckets for 15/50 mL conical centrifuge tubes</td><td>or equivalent</td></tr><tr><td>Density gradient centrifugation solution</td><td>Millipore Sigma</td><td>P1644</td><td>Percoll</td></tr><tr><td>Dimethylsulfoxide</td><td></td><td></td><td>any suitable tissue culture grade</td></tr><tr><td>Dissociation enzyme solution</td><td>StemCell Technologies</td><td>07921</td><td>ACCUMAX</td></tr><tr><td>DMEM-F12</td><td>ThermoFisher Scientific</td><td>11039021</td><td>or equivalent</td></tr><tr><td>Forceps</td><td></td><td></td><td>any suitable</td></tr><tr><td>HA hydrogel kit</td><td>ESI BIO</td><td>GS311</td><td>HyStem (Hyaluronic acid-SH and PEGDA)</td></tr><tr><td>Hanks Balanced Salt Solution</td><td>Lonza</td><td>10-527F</td><td>or equivalent</td></tr><tr><td>Heat-inactivated fetal bovine serum</td><td>Atlanta Biologicals</td><td>S11150</td><td></td></tr><tr><td>Hemocytometer</td><td>Fisher Scientific</td><td>02-671-51B Hausser BrightLine</td><td>or equivalent</td></tr><tr><td>Hoechst 33342</td><td>ThermoFisher Scientific</td><td>H1398</td><td>or equivalent</td></tr><tr><td>Image processing software</td><td>Oxford Instruments</td><td>Imaris 9.3</td><td>or equivalent</td></tr><tr><td>LIVE/DEAD Cell Viability/Cytotoxicity Kit (Calcein-AM/Ethidium Homodimer-1)</td><td>ThermoFisher Scientific</td><td>L3224</td><td>or equivalent</td></tr><tr><td>Microfluidic culture plate</td><td>Mimetas</td><td>9603-400-B</td><td>2-lane OrganoPlate</td></tr><tr><td>Microscope</td><td>Nikon</td><td>A1R</td><td>or equivalent</td></tr><tr><td>Multichannel pipette</td><td>Eppendorf</td><td>3125000036</td><td>or equivalent</td></tr><tr><td>PDX-derived tumor tissue</td><td></td><td></td><td>obtained under IRB approval for human tissue and IACUC approval for animal host</td></tr><tr><td>Penicillin-streptomycin</td><td>ThermoFisher Scientific</td><td>15140-122</td><td>or equivalent</td></tr><tr><td>Perfusion rocker</td><td>Mimetas</td><td>OrganoPlate Perfusion Rocker Mini</td><td></td></tr><tr><td>pH strips (pH 5-9)</td><td></td><td></td><td>any suitable</td></tr><tr><td>Phosphate-buffered saline solution</td><td>Lonza</td><td>17-516F</td><td>or equivalent</td></tr><tr><td>Razor blades</td><td></td><td></td><td>any suitable</td></tr><tr><td>Rotating xy-shaker</td><td>VWR</td><td>Advanced 3500 Orbital Shaker</td><td>or equivalent</td></tr><tr><td>Scalpel handle</td><td></td><td></td><td>any suitable</td></tr><tr><td>Single channel repeating pipette</td><td>Eppendorf</td><td>22260201</td><td></td></tr><tr><td>Sterile, 15mL conical centrifuge tubes</td><td></td><td></td><td>any suitable</td></tr><tr><td>Sterile, 50mL conical centrifuge tubes</td><td></td><td></td><td>any suitable</td></tr></tbody></table>

enhanced viability for ex vivo 3d hydrogel cultures of patient-derived xenografts in a perfused microfluidic platform

Patient-derived xenografts (PDX), generated when resected patient tumor tissue is engrafted directly into immunocompromised mice, remain biologically stable, thereby preserving molecular, genetic, and histological features, as well as heterogeneity of the original tumor. However, using these models to perform a multitude of experiments, including drug screening, is prohibitive both in terms of cost and time. Three-dimensional (3D) culture systems are widely viewed as platforms in which cancer cells retain their biological integrity through biochemical interactions, morphology, and architecture. Our team has extensive experience culturing PDX cells in vitro using 3D matrices composed of hyaluronic acid (HA). In order to separate mouse fibroblast stromal cells associated with PDXs, we use rotation culture, where stromal cells adhere to the surface of tissue culture-treated plates while dissociated PDX tumor cells float and self-associate into multicellular clusters. Also floating in the supernatant are single, often dead cells, which present a challenge in collecting viable PDX clusters for downstream encapsulation into hydrogels for 3D cell culture. In order to separate these single cells from live cell clusters, we have employed density step gradient centrifugation. The protocol described here allows for the depletion of non-viable single cells from the healthy population of cell clusters that will be used for further in vitro experimentation. In our studies, we incorporate the 3D cultures in microfluidic plates which allow for media perfusion during culture. After assessing the resultant cultures using a fluorescent image-based viability assay of purified versus non-purified cells, our results show that this additional separation step substantially reduced the number of non-viable cells from our cultures.

A programmable perfusion rocker was prepared in a standard water-jacked cell culture incubator, and two-lane microfluidic plates were prepared in a standard biosafety cabinet for loading (Figure 1). An MDA-PCA-118b PDX tumor was expanded in vivo, harvested when it had reached a maximum size, and dissociated as described in protocol section 2 to create a slurry suspension of cells, at approximately a single-cell state (Figure 2A). The slurry was dispensed into 6-...

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Enhanced Viability for Ex vivo 3D Hydrogel Cultures of Patient-Derived Xenografts in a Perfused Microfluidic Platform

This protocol demonstrates methods to enable extended in vitro culture of patient-derived xenografts (PDX). One step enhances overall viability of multicellular cluster cultures in 3D hydrogels, through straightforward removal of non-viable single cells. A secondary step demonstrates best practices for PDX culture in a perfused microfluidic platform.

Patient-derived xenografts (PDX), generated when resected patient tumor tissue is engrafted directly into immunocompromised mice, remain biologically ...

enhanced-viability-for-ex-vivo-3d-hydrogel-cultures-patient-derived

Research

JoVE Journal

Cancer Research

6.0K Views.  Rice University. Verbeterde kweekmethoden voor door patiënten afgeleide tumormonsters zijn van cruciaal belang omdat het veld zich afbeweet van het gebruik van conventionele cellijnen. Dit protocol maakt de in vitro cultuur van patiënt-afgeleide tumorcellen in een platform dat vatbaar is voor een hoge doorvoer en hoge inhoud screening, in tegenstelling tot in vivo kanker modellen. Het platform met PDXs stelt ons in staat om meer heterogene systemen te beoordelen die reflecteren op inheemse tumoren.Di...

Video: Enhanced Viability for Ex vivo 3D Hydrogel Cultures of Patient-Derived Xenografts in a Perfused Microfluidic Platform

Microfluidic Device for Recreating a Tumor Microenvironment in Vitro

We have developed a microfluidic device that mimics the delivery and systemic clearance of drugs to heterogeneous three-dimensional tumor tissues in vitro. Nutrients delivered by vasculature fail to reach all parts of tumors, giving rise to heterogeneous microenvironments consisting of viable, quiescent and necrotic cell types. Many cancer drugs fail to effectively penetrate and treat all types of cells because of this heterogeneity. Monolayers of cancer cells do not mimic this heterogeneity, making it difficult to test cancer drugs with a suitable in vitro model. Our microfluidic devices were fabricated out of PDMS using soft lithography. Multicellular tumor spheroids, formed by the hanging drop method, were inserted and constrained into rectangular chambers on the device and maintained with continuous medium perfusion on one side. The rectangular shape of chambers on the device created linear gradients within tissue. Fluorescent stains were used to quantify the variability in apoptosis within tissue. Tumors on the device were treated with the fluorescent chemotherapeutic drug doxorubicin, time-lapse microscopy was used to monitor its diffusion into tissue, and the effective diffusion coefficient was estimated. The hanging drop method allowed quick formation of uniform spheroids from several cancer cell lines. The device enabled growth of spheroids for up to 3 days. Cells in proximity of flowing medium were minimally apoptotic and those far from the channel were more apoptotic, thereby accurately mimicking regions in tumors adjacent to blood vessels. The estimated value of the doxorubicin diffusion coefficient agreed with a previously reported value in human breast cancer. Because the penetration and retention of drugs in solid tumors affects their efficacy, we believe that this device is an important tool in understanding the behavior of drugs, and developing new cancer therapeutics.

Microfluidic Device for Recreating a Tumor Microenvironment in Vitro

We present the procedure for fabrication and operation of a microfluidic device that recreates heterogeneous tumor microenvironments in vitro. The variability in apoptosis within tumor tissue was quantified using fluorescent stains and the effective diffusion coefficient of the chemotherapeutic drug doxorubicin into tumor tissue was evaluated.

We have developed a microfluidic device that mimics the delivery and systemic clearance of drugs to heterogeneous three-dimensional tumor tissues in ...

Bioengineering

Endothelialized Microfluidics for Studying Microvascular Interactions in Hematologic Diseases

Advances in microfabrication techniques have enabled the production of inexpensive and reproducible microfluidic systems for conducting biological and biochemical experiments at the micro- and nanoscales 1,2. In addition, microfluidics have also been specifically used to quantitatively analyze hematologic and microvascular processes, because of their ability to easily control the dynamic fluidic environment and biological conditions3-6. As such, researchers have more recently used microfluidic systems to study blood cell deformability, blood cell aggregation, microvascular blood flow, and blood cell-endothelial cell interactions6-13.However, these microfluidic systems either did not include cultured endothelial cells or were larger than the sizescale relevant to microvascular pathologic processes. A microfluidic platform with cultured endothelial cells that accurately recapitulates the cellular, physical, and hemodynamic environment of the microcirculation is needed to further our understanding of the underlying biophysical pathophysiology of hematologic diseases that involve the microvasculature.
Here, we report a method to create an "endothelialized" in vitro model of the microvasculature, using a simple, single mask microfabrication process in conjunction with standard endothelial cell culture techniques, to study pathologic biophysical microvascular interactions that occur in hematologic disease. This "microvasculature-on-a-chip" provides the researcher with a robust assay that tightly controls biological as well as biophysical conditions and is operated using a standard syringe pump and brightfield/fluorescence microscopy. Parameters such as microcirculatory hemodynamic conditions, endothelial cell type, blood cell type(s) and concentration(s), drug/inhibitory concentration etc., can all be easily controlled. As such, our microsystem provides a method to quantitatively investigate disease processes in which microvascular flow is impaired due to alterations in cell adhesion, aggregation, and deformability, a capability unavailable with existing assays.

A method to culture an endothelial cell monolayer throughout the entire inner 3D surface of a microfluidic device with microvascular-sized channels (&lt;30 &mu;m) is described. This in vitro microvasculature model enables the study of biophysical interactions between blood cells, endothelial cells, and soluble factors in hematologic diseases.

Advances in  microfabrication techniques have enabled the production of inexpensive and  reproducible microfluidic systems for conducting biological and ...

An In Vitro 3D Model and Computational Pipeline to Quantify the Vasculogenic Potential of iPSC-Derived Endothelial Progenitors

Induced pluripotent stem cells (iPSCs) are a patient-specific, proliferative cell source that can differentiate into any somatic cell type. Bipotent endothelial progenitors (EPs), which can differentiate into the cell types necessary to assemble mature, functional vasculature, have been derived from both embryonic and induced pluripotent stem cells. However, these cells have not been rigorously evaluated in three-dimensional environments, and a quantitative measure of their vasculogenic potential remains elusive. Here, the generation and isolation of iPSC-EPs via fluorescent-activated cell sorting are first outlined, followed by a description of the encapsulation and culture of iPSC-EPs in collagen hydrogels. This extracellular matrix (ECM)-mimicking microenvironment encourages a robust vasculogenic response; vascular networks form after a week of culture. The creation of a computational pipeline that utilizes open-source software to quantify this vasculogenic response is delineated. This pipeline is specifically designed to preserve the 3D architecture of the capillary plexus to robustly identify the number of branches, branching points, and the total network length with minimal user input.

Endothelial progenitors derived from induced pluripotent stem cells (iPSC-EPs) have the potential to revolutionize cardiovascular disease treatments and to enable the creation of more faithful cardiovascular disease models. Herein, the encapsulation of iPSC-EPs in three-dimensional (3D) collagen microenvironments and a quantitative analysis of these cells&#8217; vasculogenic potential are described.

Induced pluripotent stem cells (iPSCs) are a patient-specific, proliferative cell source that can differentiate into any somatic cell type. Bipotent ...

Developmental Biology

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 ...

3D Microtissues for Injectable Regenerative Therapy and High-throughput Drug Screening

To upgrade traditional 2D cell culture to 3D cell culture, we have integrated microfabrication with cryogelation technology to produce macroporous microscale cryogels (microcryogels), which can be loaded with a variety of cell types to form 3D microtissues. Herein, we present the protocol to fabricate versatile 3D microtissues and their applications in regenerative therapy and drug screening. Size and shape-controllable microcryogels can be fabricated on an array chip, which can be harvested off-chip as individual cell-loaded carriers for injectable regenerative therapy or be further assembled on-chip into 3D microtissue arrays for high-throughput drug screening. Due to the high elastic nature of these microscale cryogels, the 3D microtissues exhibit great injectability for minimally invasive cell therapy by protecting cells from mechanical shear force during injection. This ensures enhanced cell survival and therapeutic effect in the mouse limb ischemia model. Meanwhile, assembly of 3D microtissue arrays in a standard 384-multi-well format facilitates the use of common laboratory facilities and equipment, enabling high-throughput drug screening on this versatile 3D cell culture platform.

This protocol describes the fabrication of elastic 3D macroporous microcryogels by integrating microfabrication with cryogelation technology. Upon loading with cells, 3D microtissues are generated, which can be readily injected in vivo to facilitate regenerative therapy or assembled into arrays for in vitro high-throughput drug screening.

To upgrade traditional 2D cell culture to 3D cell culture, we have integrated microfabrication with cryogelation technology to produce macroporous ...

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 ...

Standardized and Scalable Assay to Study Perfused 3D Angiogenic Sprouting of iPSC-derived Endothelial Cells In Vitro

Pre-clinical drug research of vascular diseases requires in vitro models of vasculature that are amendable to high-throughput screening. However, current in vitro screening models that have sufficient throughput only have limited physiological relevance, which hinders the translation of findings from in vitro to in vivo. On the other hand, microfluidic cell culture platforms have shown unparalleled physiological relevancy in vitro, but often lack the required throughput, scalability and standardization. We demonstrate a robust platform to study angiogenesis of endothelial cells derived from human induced pluripotent stem cells (iPSC-ECs) in a physiological relevant cellular microenvironment, including perfusion and gradients. The iPSC-ECs are cultured as 40 perfused 3D microvessels against a patterned collagen-1 scaffold. Upon the application of a gradient of angiogenic factors, important hallmarks of angiogenesis can be studied, including the differentiation into tip- and stalk cell and the formation of perfusable lumen. Perfusion with fluorescent tracer dyes enables the study of permeability during and after anastomosis of the angiogenic sprouts. In conclusion, this method shows the feasibility of iPSC-derived ECs in a standardized and scalable 3D angiogenic assay that combines physiological relevant culture conditions in a platform that has the required robustness and scalability to be integrated within the drug screening infrastructure.

This method describes the culture of iPSC-derived endothelial cells as 40 perfused 3D microvessels in a standardized microfluidic platform. This platform enables the study of gradient-driven angiogenic sprouting in 3D, including anastomosis and stabilization of the angiogenic sprouts in a scalable and high-throughput manner.

Pre-clinical drug research of vascular diseases requires in vitro models of vasculature that are amendable to high-throughput screening. However, current ...

Developing 3D Organized Human Cardiac Tissue within a Microfluidic Platform

The leading cause of death worldwide persists as cardiovascular disease (CVD). However, modeling the physiological and biological complexity of the heart muscle, the myocardium, is notoriously difficult to accomplish in vitro. Mainly, obstacles lie in the need for human cardiomyocytes (CMs) that are either adult or exhibit adult-like phenotypes and can successfully replicate the myocardium&#39;s cellular complexity and intricate 3D architecture. Unfortunately, due to ethical concerns and lack of available primary patient-derived human cardiac tissue, combined with the minimal proliferation of CMs, the sourcing of viable human CMs has been a limiting step for cardiac tissue engineering. To this end, most research has transitioned toward cardiac differentiation of human induced pluripotent stem cells (hiPSCs) as the primary source of human CMs, resulting in the wide incorporation of hiPSC-CMs within in vitro assays for cardiac tissue modeling.
Here in this work, we demonstrate a protocol for developing a 3D mature stem cell-derived human cardiac tissue within a microfluidic device. We specifically explain and visually demonstrate the production of a 3D in vitro anisotropic cardiac tissue-on-a-chip model from hiPSC-derived CMs. We primarily describe a purification protocol to select for CMs, the co-culture of cells with a defined ratio via mixing CMs with human CFs (hCFs), and suspension of this co-culture within the collagen-based hydrogel. We further demonstrate the injection of the cell-laden hydrogel within our well-defined microfluidic device, embedded with staggered elliptical microposts that serve as surface topography to induce a high degree of alignment of the surrounding cells and the hydrogel matrix, mimicking the architecture of the native myocardium. We envision that the proposed 3D anisotropic cardiac tissue-on-chip model is suitable for fundamental biology studies, disease modeling, and, through its use as a screening tool, pharmaceutical testing.

The goal of this protocol is to explain and demonstrate the development of a three-dimensional (3D) microfluidic model of highly aligned human cardiac tissue, composed of stem cell-derived cardiomyocytes co-cultured with cardiac fibroblasts (CFs) within a biomimetic, collagen-based hydrogel, for applications in cardiac tissue engineering, drug screening, and disease modeling.

The leading cause of death worldwide persists as cardiovascular disease (CVD). However, modeling the physiological and biological complexity of the heart ...

Hydrogel Arrays Enable Increased Throughput for Screening Effects of Matrix Components and Therapeutics in 3D Tumor Models

Cell-matrix interactions mediate complex physiological processes through biochemical, mechanical, and geometrical cues, influencing pathological changes and therapeutic responses. Accounting for matrix effects earlier in the drug development pipeline is expected to increase the likelihood of clinical success of novel therapeutics. Biomaterial-based strategies recapitulating specific tissue microenvironments in 3D cell culture exist but integrating these with the 2D culture methods primarily used for drug screening has been challenging. Thus, the protocol presented here details the development of methods for 3D culture within miniaturized biomaterial matrices in a multi-well plate format to facilitate integration with existing drug screening pipelines and conventional assays for cell viability. Since the matrix features critical for preserving clinically relevant phenotypes in cultured cells are expected to be highly tissue- and disease-specific, combinatorial screening of matrix parameters will be necessary to identify appropriate conditions for specific applications. The methods described here use a miniaturized culture format to assess cancer cell responses to orthogonal variation of matrix mechanics and ligand presentation. Specifically, this study demonstrates the use of this platform to investigate the effects of matrix parameters on the responses of patient-derived glioblastoma (GBM) cells to chemotherapy.

The present protocol describes an experimental platform to assess the effects of mechanical and biochemical cues on chemotherapeutic responses of patient-derived glioblastoma cells in 3D matrix-mimetic cultures using a custom-made UV illumination device facilitating high-throughput photocrosslinking of hydrogels with tunable mechanical features.

Cell-matrix interactions mediate complex physiological processes through biochemical, mechanical, and geometrical cues, influencing pathological changes ...

3D Co-Culture Assay: Vascularized Osteogenic Niches for Cancer Drug Screening

Simple Establishment of a Vascularized Osteogenic Bone Marrow Niche Using Pre-Cast Poly(ethylene Glycol) (PEG) Hydrogels in an Imaging Microplate

The bone and bone marrow are highly vascularized and structurally complex organs, and are sites for cancer and metastasis formation. In vitro models recapitulating bone- and bone marrow-specific functions, including vascularization, that are compatible with drug screening are highly desirable. Such models can bridge the gap between simplistic, structurally irrelevant two-dimensional (2D) in vitro models and the more expensive, ethically challenging in vivo models. This article describes a controllable three-dimensional (3D) co-culture assay based on engineered poly(ethylene glycol) (PEG) matrices for the generation of vascularized, osteogenic bone-marrow niches. The PEG matrix design allows the development of 3D cell cultures through a simple cell seeding step requiring no encapsulation, thus enabling the development of complex co-culture systems. Furthermore, the matrices are transparent and pre-cast onto glass-bottom 96-well imaging plates, rendering the system suitable for microscopy. For the assay described here, human bone marrow-derived mesenchymal stromal cells (hBM-MSCs) are cultured first until a sufficiently developed 3D cell network is formed. Subsequently, GFP-expressing human umbilical vein endothelial cells (HUVECs) are added. The culture development is followed by bright-field and fluorescence microscopy. The presence of the hBM-MSC network supports the formation of vascular-like structures that otherwise&#160;would not form and that remain stable for at least 7 days. The extent of vascular-like network formation can easily be quantified. This model can be tuned toward an osteogenic bone-marrow niche by supplementing the culture medium with bone morphogenetic protein 2 (BMP-2), which promotes the osteogenic differentiation of the hBM-MSCs, as assessed by increased alkaline phosphatase (ALP) activity at day 4 and day 7 of co-culture. This cellular model can be used as a platform for culturing various cancer cells and studying how they interact with bone- and bone marrow-specific vascular niches. Moreover, it is suitable for automation and high-content analyses, meaning it would enable cancer drug screening under highly reproducible culture conditions.

Author Spotlight: Simple Establishment of a Vascularized Osteogenic Bone Marrow Niche Using Pre-Cast Poly(Ethylene Glycol) (PEG) Hydrogels in an Imaging Microplate  

An in vitro model of the bone marrow vascular niche is established by seeding mesenchymal and endothelial cells onto pre-cast 3D PEG hydrogels. The endothelial networks, ECM components, and ALP activity of the niches vary depending on the growth factor used. The platform can be used for advanced cancer models.

The bone and bone marrow are highly vascularized and structurally complex organs, and are sites for cancer and metastasis formation. In vitro models ...

Enhanced Viability for Ex vivo 3D Hydrogel Cultures of Patient-Derived Xenografts in a Perfused Microfluidic Platform

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