This work was supported by Hong Kong Research Grant Council (grants 17122014, C1013-15G, 719813E, and 17304514). A. S. T. Wong is a recipient of the Croucher Senior Research Fellowship.

1. Microfluidic Device Design and Fabrication
<ol>
	<li>Microfluidic master design
	<ol>
		<li>Design and draw the microfluidic channel pattern with any computer-aided design (CAD) software. 
		NOTE: Normally, the CAD drawing can be sent to a photomask company to produce the photomask. The microfluidic design consists of three identical parallel channels, each with the following dimensions: 4 mm &#215; 25 mm &#215; 250 &#181;m (width &#215; length &#215; height) and set 2 mm apart. Both channel ends were designed with 127&#176; angles to facilitate liquid entrance and exit (Figure 1B). The channel used in this protocol was reported in a previous publication13.</li>
		<li>Wash the silicon wafer with ultrasonication (500 W/42 kHz) in acetone, isopropanol, and deionized water for 15 min each. Dry the wafer with pressurized nitrogen gas. Heat the silicon wafer on a hot plate at 250 &#176;C for 15 min to completely dry it.</li>
		<li>Spin-coat a layer of SU-8 2075 photoresist (125 &#181;m-thick) onto the silicon wafer with a spinning speed of 1,800 rpm. Bake the photoresist-coated wafer directly on a hot plate at 65 &#176;C and then at 95 &#176;C, to remove excess solvent, for 5 and 25 min, respectively (according to the SU-8 product manual).</li>
		<li>Repeat step 3 and spin-coat another layer of SU-8 2075 photoresist to obtain a final thickness of 250 &#181;m. Bake the photoresist-coated wafer directly on a hot plate at 65 &#176;C for 7 min and then at 95 &#176;C for 30 min. Slowly cool the wafer to room temperature. Do not cool with pressurized air.</li>
		<li>Align and put the photomask directly on top of the wafer. Expose it to UV light for 20 s to crosslink the pattern.</li>
		<li>Bake the wafer with photoresist at 65 &#176;C and 95 &#176;C on a hotplate for 5 and 12 min, respectively, to remove the solvent for polymerization (according to the SU-8 product manual). Afterwards, cool the wafer to room temperature.</li>
		<li>Remove the non-crosslinked photoresist by immersing the wafer in SU-8 developer for 25 min. Rinse the wafer with isopropanol and dry it with pressured air. 
		NOTE: If a milky solution was observed when rinsing with isopropanol, indicating incomplete development, immerse the wafer back into SU-8 developer for a longer development time.</li>
		<li>Place the wafer on a hot plate at 180 &#176;C for 10 min to heal the pattern from potential cracks. Turn off the hot plate and slowly cool down the wafer to room temperature by leaving it on the hot plate.</li>
		<li>Add a few drops of trichloro (1H, 1H, 2H, 2H-perfluorooctyl) silane in a vial cap. Put the vial cap in the vacuum desiccator. Place the wafer next to the vial cap to silanize the wafer for 10 min.</li>
	</ol>
	</li>
	<li>PDMS chip fabrication
	<ol>
		<li>Wrap the wafer with aluminum foil to create a holder.</li>
		<li>Mix the polydimethylsiloxane (PDMS) base and curing agent at a 10:1 mass ratio with a centrifugal mixer under the mixing and degas function. Pour the PDMS mixture slowly onto the wafer to a height of around 5 mm. Degas the PDMS under vacuum for 40 min. Cure the PDMS in an oven at 65 &#176;C for 2 h.</li>
		<li>Carefully peel the PDMS slab off the master and trim the PDMS along the channels until it is the size of a glass slide. Punch the PDMS with a 1-mm biopsy punch to create the inlets and outlets of the device.</li>
		<li>Clean the PDMS surface with pressurized air and adhesive tape to remove dust and PDMS debris.</li>
		<li>Place the PDMS slab, with the channel side facing upwards, into a plasma cleaner. Place a clean glass slide into the plasma cleaner alongside the PDMS slab.</li>
		<li>Treat the PDMS and glass slide under air plasma at high RF level for 1 min. Bind the PDMS to the glass slide to create an irreversible covalent bond.</li>
		<li>Place the PDMS chip on a hot plate at 150 &#176;C for 1 h to enhance the bonding strength and to return the hydrophobicity of the PDMS before use.</li>
	</ol>
	</li>
</ol>
2. Seeding the Microfluidic Chip with Primary Peritoneal Mesothelial Cells (HPMCs)
<ol>
	<li>Fibronectin coating of microfluidic channels
	<ol>
		<li>Sterilize the microfluidic channel with 30 &#181;L of 75% ethanol. Remove the ethanol and rinse twice with 30 &#181;L of sterilized phosphate-buffered saline (PBS) using a pipette tip. Thoroughly aspirate PBS in the channels. 
		NOTE: To minimize contamination, the microfluidic cell culture should be carefully conducted under fully aseptic conditions through the use of a laminar flow hood.</li>
		<li>Prepare a 10 &#181;g/mL fibronectin solution in serum-free M199:MCDB105 (1:1) medium. Pipette the solution up and down, with special care to avoid bubble formation. Do not vortex.</li>
		<li>Slowly fill up each channel with 30 &#181;L of fibronectin solution. Seal the channels with tape and incubate the chip overnight at 4 &#176;C.</li>
	</ol>
	</li>
	<li>Seeding HPMCs into the microfluidic channels
	<ol>
		<li>Culture HPMCs in 10% fetal bovine serum (FBS) culture medium (M199:MCDB105 supplemented with 10% FBS and 100 U/mL penicillin/streptomycin) under 5% CO2 at 37 &#176;C until 80% confluence.</li>
		<li>Rinse the HPMCs with 3 mL of PBS and trypsinize with 2 mL of 0.05% trypsin/0.01% EDTA (TE) solution for 30 s. Neutralize with 4 mL of 10% FBS culture medium and spin down the cells at 1,000 x g for 5 min.</li>
		<li>Resuspend the HPMCs in 2 mL of 10% FBS culture medium. Count the cell number with a hemocytometer and adjust the cell concentration to 3.5&#215;106/mL.</li>
		<li>Warm the fibronectin-coated microfluidic chip in a 37 &#176;C incubator for 5 min before use. Completely aspirate the fibronectin solution.</li>
		<li>Slowly pipette 30 &#181;L of HPMC suspension into each channel. Seal the channels with tape and place the devices in the CO2 incubator overnight.</li>
	</ol>
	</li>
</ol>
3. Ovarian Cancer Spheroid Formation and Co-culture
<ol>
	<li>Ovarian cancer spheroid formation
	<ol>
		<li>Maintain the human epithelial ovarian cancer cell line SKOV-3 in 5% FBS culture medium (M199:MCDB105 with 5% FBS and 100 U/mL penicillin/streptomycin) under 5% CO2 at 37 &#176;C before the experiments.</li>
		<li>Dissolve 0.5% agarose in sterilized water by boiling. Dispense 50 &#181;L of agarose solution into each well of 96-well plates. Leave the plates in the hood for 20 min to allow the agarose to solidify; solidified agarose coating the wells can prevent cell adhesion and indicates that the plates are ready for use.</li>
		<li>Rinse the SKOV-3 cells with 3 mL of PBS. Trypsinize with 2 mL of TE solution for 3 min. Neutralize with 4 mL of 5% FBS culture medium and spin down the cells at 1,000 x g for 5 min.</li>
		<li>Count the SKOV-3 cell number with a hemocytometer and resuspend the cells in 5% FBS culture medium at a density of 5,000 cells/mL. Transfer 200 &#181;L of cell suspension into each agarose-coated well and culture under 5% CO2 at 37 &#176;C for 48 h.</li>
	</ol>
	</li>
	<li>Cancer spheroid loading
	<ol>
		<li>Transfer cancer spheroids to a 50-mL centrifuge tube pre-wetted with 1 mL of serum-free medium and spin down at 750 x g for 5 min. 
		NOTE: Pre-wetted pipette tips and centrifuge tubes can prevent undesired spheroid attachment onto the centrifuge tubes.</li>
		<li>Prepare 2.5 &#181;g/mL of 5-chloromethylfluorescein diacetate (CMFDA) solution in serum-free M199:MCDB105 medium. Resuspend cancer spheroids in 1 mL of CMFDA solution and incubate at 37 &#176;C for 30 min. Centrifuge at 750 x g for 5 min.</li>
		<li>Rinse the cancer spheroids with 2 mL of serum-free medium and centrifuge at 750 x g for 5 min. Repeat this step twice to remove the remaining fluorescent dye from the medium. 
		NOTE: The fluorescent signal can be retained in spheroids for more than 96 h.</li>
		<li>Resuspend the cancer spheroids in serum-free M199:MCDB105 medium and count the spheroid number with a hemocytometer. Adjust the concentration to 2,000 spheroids/mL with serum-free medium.</li>
		<li>Prior to spheroid loading, slowly inject 100 &#181;L of serum-free M199:MCDB105 medium to wash away the non-adhered mesothelial cells inside the channels. 
		NOTE: A fast injection, aspiration, or bubble formation will lead to the detachment of the mesothelial monolayer.</li>
		<li>Load 30 &#181;L of fluorescent-labeled spheroid suspension into each channel. Place the microfluidic chip into the CO2 incubator.</li>
	</ol>
	</li>
	<li>Perfusion platform assembly and co-culture
	<ol>
		<li>Attach a syringe loaded with fresh, serum-free M199:MCDB105 medium to a 1 m-long tube.</li>
		<li>Place the syringe on the syringe pump and secure the plunger and the body of the syringe. Install the syringe pump in a horizontal position at the same height as the microfluidic chip inside the incubator. 
		NOTE: The number and size of syringes depend on the number of channels used and the duration of the experiment.</li>
		<li>Quickly infuse the whole tube with medium by pressing and holding the fast forward button until the medium overflows the tubing.</li>
		<li>Run the injection at the desired flow rate. Estimate the wall shear stress using the following equation: 
		<img alt="Equation 1" src="/files/ftp_upload/55337/eq1.jpg" /> 
		where &#964; represents the wall shear stress inside the chamber, &#490; = flow rate, &#956; = viscosity (0.01 g cms-1); h = flow chamber height, and w = flow chamber width.</li>
		<li>Connect the tubing to the inlet of the channel. Place the excess inlet tubing in the incubator to ensure that the medium is warmed before injected into the channels. To facilitate the connection between the tubing and the channel, appropriate connectors can be chosen. NOTE: Beware not to introduce any bubble to the syringe, tubing, and connections.</li>
		<li>Connect the outlet of the channel to a 50-mL conical tube to collect the outflow medium with a short output tube. 
		NOTE: The complete dynamic co-culture system is shown in Figure 1D.</li>
	</ol>
	</li>
</ol>
4. Observation of HPMCs and Cancer Spheroids after Perfusion
<ol>
	<li>Observe the HPMCs and cancer spheroids with a bright-field or fluorescence microscope and take pictures after 1 h of perfusion.</li>
</ol>

<ol>
	<li>Jemal, A., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Global+cancer+statistics.">Global cancer statistics.</a> CA: Cancer J. Clin. 61, 69-90 (2011).</li><li>Burleson, K. M., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Ovarian+carcinoma+ascites+spheroids+adhere+to+extracellular+matrix+components+and+mesothelial+cell+monolayers.">Ovarian carcinoma ascites spheroids adhere to extracellular matrix components and mesothelial cell monolayers.</a> Gynecol. Oncol. 93, 170-181 (2004).</li><li>Chau, W. K., Ip, C. K., Mak, A. S., Lai, H. C., Wong, A. S. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=c-Kit+mediates+chemoresistance+and+tumor-initiating+capacity+of+ovarian+cancer+cells+through+activation+of+Wnt/beta-catenin-ATP-binding+cassette+G2+signaling.">c-Kit mediates chemoresistance and tumor-initiating capacity of ovarian cancer cells through activation of Wnt/beta-catenin-ATP-binding cassette G2 signaling.</a> Oncogene. 32, 2767-2781 (2013).</li><li>Zhang, S., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Identification+and+characterization+of+ovarian+cancer-initiating+cells+from+primary+human+tumors.">Identification and characterization of ovarian cancer-initiating cells from primary human tumors.</a> Cancer Res. 68, 4311-4320 (2008).</li><li>Tang, M. K. S., Zhou, H. Y., Yam, J. W. P., Wong, A. S. T. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=c-Met+overexpression+contributes+to+the+acquired+apoptotic+resistance+of+nonadherent+ovarian+cancer+cells+through+a+cross+talk+mediated+by+phosphatidylinositol+3-kinase+and+extracellular+signal-regulated+kinase+1/2.">c-Met overexpression contributes to the acquired apoptotic resistance of nonadherent ovarian cancer cells through a cross talk mediated by phosphatidylinositol 3-kinase and extracellular signal-regulated kinase 1/2.</a> Neoplasia. 12, 128-144 (2010).</li><li>Ksiazek, K., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Senescent+peritoneal+mesothelial+cells+promote+ovarian+cancer+cell+adhesion:+the+role+of+oxidative+stress-induced+fibronectin.">Senescent peritoneal mesothelial cells promote ovarian cancer cell adhesion: the role of oxidative stress-induced fibronectin.</a> Am. J. Pathol. 174, 1230-1240 (2009).</li><li>Hafter, R., Klaubert, W., Gollwitzer, R., Vonhugo, R., Graeff, H. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Crosslinked+Fibrin+Derivatives+and+Fibronectin+in+Ascitic+Fluid+from+Patients+with+Ovarian-Cancer+Compared+to+Ascitic+Fluid+in+Liver-Cirrhosis.">Crosslinked Fibrin Derivatives and Fibronectin in Ascitic Fluid from Patients with Ovarian-Cancer Compared to Ascitic Fluid in Liver-Cirrhosis.</a> Thromb Res. 35, 53-64 (1984).</li><li>Kenny, H. A., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Mesothelial+cells+promote+early+ovarian+cancer+metastasis+through+fibronectin+secretion.">Mesothelial cells promote early ovarian cancer metastasis through fibronectin secretion.</a> J. Clin. Invest. 124, 4614-4628 (2014).</li><li>Jain, R. K. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Normalization+of+tumor+vasculature:+An+emerging+concept+in+antiangiogenic+therapy.">Normalization of tumor vasculature: An emerging concept in antiangiogenic therapy.</a> Science. 307, 58-62 (2005).</li><li>Chang, S. F., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Tumor+cell+cycle+arrest+induced+by+shear+stress:+Roles+of+integrins+and+Smad.">Tumor cell cycle arrest induced by shear stress: Roles of integrins and Smad.</a> Proc. Natl. Acad. Sci. USA. 105, 3927-3932 (2008).</li><li>Rutkowski, J. M., Swartz, M. A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=A+driving+force+for+change:+interstitial+flow+as+a+morphoregulator.">A driving force for change: interstitial flow as a morphoregulator.</a> Trends Cell Biol. 17, 44-50 (2007).</li><li>Rizvi, I., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Flow+induces+epithelial-mesenchymal+transition,+cellular+heterogeneity+and+biomarker+modulation+in+3D+ovarian+cancer+nodules.">Flow induces epithelial-mesenchymal transition, cellular heterogeneity and biomarker modulation in 3D ovarian cancer nodules.</a> Proc. Natl. Acad. Sci. USA. 110, E1974-E1983 (2013).</li><li>Ip, C. K., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Stemness+and+chemoresistance+in+epithelial+ovarian+carcinoma+cells+under+shear+stress.">Stemness and chemoresistance in epithelial ovarian carcinoma cells under shear stress.</a> Sci. Rep. 6, 26788 (2016).</li><li>Avraham-Chakim, L., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Fluid-flow+induced+wall+shear+stress+and+epithelial+ovarian+cancer+peritoneal+spreading.">Fluid-flow induced wall shear stress and epithelial ovarian cancer peritoneal spreading.</a> PloS one. 8, e60965 (2013).</li><li>Burkhalter, R. J., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Peritoneal+mechanobiology+and+metastatic+success+in+epithelial+ovarian+cancer.">Peritoneal mechanobiology and metastatic success in epithelial ovarian cancer.</a> Faseb Journal. 26, (2012).</li><li>Lane, W. O., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Parallel-plate+flow+chamber+and+continuous+flow+circuit+to+evaluate+endothelial+progenitor+cells+under+laminar+flow+shear+stress.">Parallel-plate flow chamber and continuous flow circuit to evaluate endothelial progenitor cells under laminar flow shear stress.</a> Journal of visualized experiments : JoVE. , (2012).</li><li>Botta, G. P., Manley, P., Miller, S., Lelkes, P. I. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Real-time+assessment+of+three-dimensional+cell+aggregation+in+rotating+wall+vessel+bioreactors+in+vitro.">Real-time assessment of three-dimensional cell aggregation in rotating wall vessel bioreactors in vitro.</a> Nat. Protoc. 1, 2116-2127 (2006).</li><li>Ismadi, M. Z., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Flow+characterization+of+a+spinner+flask+for+induced+pluripotent+stem+cell+culture+application.">Flow characterization of a spinner flask for induced pluripotent stem cell culture application.</a> PloS one. 9, e106493 (2014).</li><li>Yu, W., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=A+microfluidic-based+multi-shear+device+for+investigating+the+effects+of+low+fluid-induced+stresses+on+osteoblasts.">A microfluidic-based multi-shear device for investigating the effects of low fluid-induced stresses on osteoblasts.</a> PloS one. 9. 9, e89966 (2014).</li></ol>

The authors have nothing to disclose.

This assay offers a flexible and physiologically relevant model that can be incorporated with various biochemical and cell-based assays, including, but not limited to, adhesion assays, mesothelial clearance assays, and drug screening. It can be applied to the evaluation of the effect of the intraperitoneal microenvironment on cancer progression. However, several experimental conditions might need to be optimized, depending on the aims of the project (e.g., the number of HPMCs and cancer spheroids seeded per channel, the co-culture time, etc.). Other cell types in the intraperitoneal microenvironment, such as fibroblasts, endothelial cells, or immune cells, can also be included in the system in order to study the essential interactions between ovarian cancer spheroids and nearby cells. Other ECM components, such as laminin, vitronectin, or collagen, can also be used. One weakness of this protocol that may need to be considered is that, while the coupling of nutrient replenishing and shear stress is reminiscent of that seen in vivo, there is no separate control of nutrient replenishing and shear stress in our current system. Moreover, techniques such as on-chip trypsinization combined with fluorescence-activated cell sorting will be needed to separately collect ovarian cancer cells and HPMCs for further molecular studies about different types of cells.
This technique exhibits several advantages when compared with traditional approaches. First, 3D co-culture better mimics the in vivo intercellular communication between ovarian cancer cells and mesothelial cells. Second, cells are cultured under fluidic flow but are not in static condition, generating a cellular response that is potentially representative of the in vivo situation12,13. Furthermore, the fluid flow can be precisely controlled, mimicking the hydrodynamic conditions at different stages of tumor progression.
In summary, the dynamic 3D ovarian cancer-mesothelium co-culture model presented here provides a flexible framework for modeling tumor behavior in the peritoneal cavity. With this model, the interaction between cancer cells and mesothelial cells under a dynamic hydrodynamic condition can be comprehensively analyzed. It can greatly advance our understanding of the underlying mechanisms of metastatic progression and might also facilitate therapeutic development.

Ovarian cancer is the most lethal gynecological cancer and is characterized by widespread peritoneal dissemination and the formation of malignant ascites1. This extensive peritoneal metastasis represents a major clinical challenge and is associated with poor clinical outcomes. Unlike most solid carcinomas that metastasize via the blood, ovarian cancer primarily disseminates within the peritoneal cavity. Tumor cells exist as multicellular aggregates/spheroids during the process of metastasis2. The fact that suspension culture can enrich ovarian cancer stem/tumor-initiating cells further suggests that these spheroids may be associated with both tumor aggressiveness and enhanced chemoresistance3,4. There are differences in drug response between 2D and 3D cultures, which presumably have different molecular mechanisms5.
The essential interaction with the mesothelium constructs the primary microenvironment for ovarian tumor progression. These mesothelial cells lie on an extracellular matrix (ECM), where fibronectin is a ubiquitous constituent. A link between the increased expression of mesothelial cell-derived fibronectin and tumor progression has been shown. Fibronectin is abundantly present in malignant ascites6,7. Ovarian cancer cells are also able to induce the secretion of fibronectin from mesothelial cells in order to promote early ovarian cancer metastasis8.
Emerging evidence shows that mechanical stimuli, including shear stress, can modulate cell morphology, gene expression, and, thus, the phenotypes of tumor cells9,10,11. As malignant ascites develop and accumulate during tumor progression, ovarian tumor cells are exposed to fluid flow and the resulting shear stress. A number of groups, ours included, have shown the impact of shear stress on ovarian cancer progression, including cytoskeleton modifications, epithelial-to-mesenchymal transitions, and cancer stemness12,13,14,15. Thus, a physiologically relevant microenvironment is important for the investigation of tumor peritoneal metastasis. However, current in vitro hydrodynamic culture systems have limitations on mimicking and controlling a constant, low, physiologically relevant shear stress16,17,18,19. Conventional in vitro approaches focusing on either the cellular or mechanical environment are still limited in mimicking the complexity of the intraperitoneal microenvironment with proper physiological relevance.
Here, in order to engineer a new model of the peritoneum to overcome the limitations of conventional strategies and to advance the study of the intraperitoneal compartment in cancer metastasis, a 3D microfluidic-based platform with controlled fluid flow was designed. In this model, ovarian cancer spheroids were co-cultured with primary human peritoneal mesothelial cells in microfluidic chips under continuous fluidic flow (Figure1A). The mesothelial cells were plated on fibronectin. Non-adherent ovarian cancer spheroids were seeded into microfluidic channels with continuous flow medium perfused by a syringe pump. Both the 3D environment and the dynamic mechanical forces are very important factors of the metastatic cascade. This platform can be used for investigating the intraperitoneal microenvironment in terms of complex cellular and co-culture interactions, as well as with regard to dynamic mechanical cues.

<table>
	<tbody>
		<tr>
			<td>Silicon wafer</td>
			<td>University wafer</td>
			<td>#1196</td>
			<td>100mm</td>
		</tr>
		<tr>
			<td>SU-8 2075 photoresist&#160;</td>
			<td>Microchem</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>SU-8 developer&#160;</td>
			<td>Microchem</td>
			<td>108-65-6</td>
			<td></td>
		</tr>
		<tr>
			<td>Trichloro (1H, 1H, 2H, 2H-perfluorooctyl) silane</td>
			<td>Sigma</td>
			<td>448931</td>
			<td></td>
		</tr>
		<tr>
			<td>Sylgard 184</td>
			<td>Dow Corning</td>
			<td>1673921</td>
			<td>Polydimethylsiloxane (PDMS) + curing agent kit</td>
		</tr>
		<tr>
			<td>Biopsy punch&#160;</td>
			<td>Miltex</td>
			<td>33-31AA</td>
			<td>1 mm diameter</td>
		</tr>
		<tr>
			<td>Plasma cleaner</td>
			<td>Harrick Plasma</td>
			<td>PDC-002</td>
			<td></td>
		</tr>
		<tr>
			<td>Polyethylene tubing</td>
			<td>SCI</td>
			<td>BB31695-PE/5</td>
			<td>0.86mm (inner diameter)</td>
		</tr>
		<tr>
			<td>Syringe</td>
			<td>Terumo</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Syringe pump</td>
			<td>Longer precision pump&#160;</td>
			<td>&#160;LSP01-2A</td>
			<td></td>
		</tr>
		<tr>
			<td>Medium 199</td>
			<td>Invitrogen</td>
			<td>31100-035</td>
			<td>Add 2.2g/L sodium bicarbonate</td>
		</tr>
		<tr>
			<td>MCDB 105 Medium</td>
			<td>Sigma</td>
			<td>M6395</td>
			<td></td>
		</tr>
		<tr>
			<td>Fetal bovine serum (FBS)</td>
			<td>Hyclone</td>
			<td>SH30068.02</td>
			<td></td>
		</tr>
		<tr>
			<td>Penicillin/streptomycin&#160;</td>
			<td>Invitrogen</td>
			<td>15070-063</td>
			<td></td>
		</tr>
		<tr>
			<td>Trypsin EDTA solution&#160;</td>
			<td>Gibco</td>
			<td>25300-054</td>
			<td>0.05% Trypsin -0.01% EDTA, phenol red</td>
		</tr>
		<tr>
			<td>Fibronectin human</td>
			<td>BD</td>
			<td>354008</td>
			<td></td>
		</tr>
		<tr>
			<td>Agarose&#160;</td>
			<td>Invitrogen</td>
			<td>15510-027</td>
			<td></td>
		</tr>
		<tr>
			<td>5-chloromethylfluorescein diacetate</td>
			<td>Life technologies</td>
			<td>C7025</td>
			<td>Green CMFDA</td>
		</tr>
		<tr>
			<td>CO2 incubator</td>
			<td>SANYO</td>
			<td>MCO-18AIC</td>
			<td></td>
		</tr>
		<tr>
			<td>Centrifuge</td>
			<td>Hitachi</td>
			<td>CT15RE</td>
			<td></td>
		</tr>
		<tr>
			<td>Fluorescent microscope</td>
			<td>Nikon</td>
			<td></td>
			<td>Model: 80i or ECLIPSE Ti; software: SPOT</td>
		</tr>
		<tr>
			<td>SKOV-3&#160;</td>
			<td></td>
			<td></td>
			<td>Gift from Dr. N Auersperg (University of British Columbia)</td>
		</tr>
	</tbody>
</table>

modeling ovarian cancer multicellular spheroid behavior in a dynamic 3d peritoneal microdevice

Ovarian cancer is characterized by extensive peritoneal metastasis, with tumor spheres commonly found in the malignant ascites. This is associated with poor clinical outcomes and currently lacks effective treatment. Both the three-dimensional (3D) environment and the dynamic mechanical forces are very important factors in this metastatic cascade. However, traditional cell cultures fail to recapitulate this natural tumor microenvironment. Thus, in vivo-like models that can emulate the intraperitoneal environment are of obvious importance. In this study, a new microfluidic platform of the peritoneum was set up to mimic the situation of ovarian cancer spheroids in the peritoneal cavity during metastasis. Ovarian cancer spheroids generated under a non-adherent condition were cultured in microfluidic channels coated with peritoneal mesothelial cells subjected to physiologically relevant shear stress. In summary, this dynamic 3D ovarian cancer-mesothelium microfluidic platform can provide new knowledge on basic cancer biology and serve as a platform for potential drug screening and development.

Using this protocol, a microfluidic platform was set up to model ovarian cancer spheroids with mesothelial cells under hydrodynamic conditions. Primary human peritoneal mesothelial cells were cultured in the microdevice for 16 h and observed under a bright-field microscope. As shown in Figure 2A, the channel bottom was successfully covered with a monolayer of HPMCs. It is important to note that bubble formation during fibronectin or HPMC patterning will lead to channel coating failure. Through non-adherent suspension culture, multicellular spheroids with diameters of approximately 100 &#181;m, which are similar in size to those found in patients&#39; ascites, were generated and labeled with green fluorescent dye, CMFDA. The spheroids were then transferred to HPMC-coated microfluidic channels and remained in suspension. Cell morphologies were observed under the microscope and images were captured (Figure 2B). The fluorescent-labeled cancer spheroids can be easily differentiated from the mesothelial cells under a fluorescence microscope (Figure 2C).
<img alt="Figure 1" src="/files/ftp_upload/55337/55337fig1.jpg" /> 
Figure 1. A Dynamic 3D Peritoneal Microdevice for Modeling Ovarian Cancer Multicellular Spheroid Behaviors. (A) Schematic diagram of a microfluidic chip used to co-culture ovarian cancer spheroids with HPMCs under flow condition. (B) Schematic diagram showing the design of the microfluidic channels. (C) Photograph of a microfluidic chip used in the protocol, with its channels infused with dye solution for clear visualization. Bar = 1 cm. (D) Photograph showing the experimental setup. <a href="https://cloudflare-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/55337/55337fig1large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 2" src="/files/ftp_upload/55337/55337fig2.jpg" /> 
Figure 2. Images of the Ovarian Cancer Spheroid-mesothelial Cell Dynamic Co-culture Model. (A) Image of the HPMC monolayer grown in the microfluidic channel. (B) Images of ovarian cancer spheroids in the microfluidic channel coated with peritoneal mesothelial cells under phase contrast microscopy (left panel) or (C) fluorescence microscopy (Green, CMFDA; right panel). Bar = 100 &#181;m. <a href="https://cloudflare-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/55337/55337fig2large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>

Watch this Scientific Journal Video about Modeling Ovarian Cancer Multicellular Spheroid Behavior in a Dynamic 3D Peritoneal Microdevice at JoVE.com

Modeling Ovarian Cancer Multicellular Spheroid Behavior in a Dynamic 3D Peritoneal Microdevice

To study ovarian tumor progression in a physiologically relevant model, multicellular spheroids were cultured in a microdevice under simulated fluid flow. This dynamic 3D model emulates the intraperitoneal environment with the cellular and mechanical components where ovarian cancer metastasis occurs.

Ovarian cancer is characterized by extensive peritoneal metastasis, with tumor spheres commonly found in the malignant ascites. This is associated with ...

modeling-ovarian-cancer-multicellular-spheroid-behavior-dynamic-3d

Universidad San Sebastian

Research

JoVE Journal

Bioengineering

8.6K Views.  University of Hong Kong. To study ovarian tumor progression in a physiologically relevant model, multicellular spheroids were cultured in a microdevice under simulated fluid flow. This dynamic 3D model emulates the intraperitoneal environment with the cellular and mechanical components where ovarian cancer metastasis occurs.

Video: Modeling Ovarian Cancer Multicellular Spheroid Behavior in a Dynamic 3D Peritoneal Microdevice

A Method for Ovarian Follicle Encapsulation and Culture in a Proteolytically Degradable 3 Dimensional System

The ovarian follicle is the functional unit of the ovary that secretes sex hormones and supports oocyte maturation. In vitro follicle techniques provide a tool to model follicle development in order to investigate basic biology, and are further being developed as a technique to preserve fertility in the clinic1-4. Our in vitro culture system employs hydrogels in order to mimic the native ovarian environment by maintaining the 3D follicular architecture, cell-cell interactions and paracrine signaling that direct follicle development 5. Previously, follicles were successfully cultured in alginate, an inert algae-derived polysaccharide that undergoes gelation with calcium ions6-8. Alginate hydrogels formed at a concentration of 0.25% w/v were the most permissive for follicle culture, and retained the highest developmental competence 9. Alginate hydrogels are not degradable, thus an increase in the follicle diameter results in a compressive force on the follicle that can impact follicle growth10. We subsequently developed a culture system based on a fibrin-alginate interpenetrating network (FA-IPN), in which a mixture of fibrin and alginate are gelled simultaneously. This combination provides a dynamic mechanical environment because both components contribute to matrix rigidity initially; however, proteases secreted by the growing follicle degrade fibrin in the matrix leaving only alginate to provide support. With the IPN, the alginate content can be reduced below 0.25%, which is not possible with alginate alone 5. Thus, as the follicle expands, it will experience a reduced compressive force due to the reduced solids content. Herein, we describe an encapsulation method and an in vitro culture system for ovarian follicles within a FA-IPN. The dynamic mechanical environment mimics the natural ovarian environment in which small follicles reside in a rigid cortex and move to a more permissive medulla as they increase in size11. The degradable component may be particularly critical for clinical translation in order to support the greater than 106-fold increase in volume that human follicles normally undergo in vivo .

A new method for ovarian follicle encapsulation in a 3D fibrin-alginate interpenetrating network is described. This system combines structural support with proteolytic degradation to support the development of immature follicles to produce mature oocytes. This method may be applied to culture cell aggregates to maintain cell-cell contacts without limiting expansion.

The ovarian follicle is the functional unit of the ovary that secretes sex hormones and supports oocyte maturation. In vitro  follicle techniques provide ...

Cellular Lipid Extraction for Targeted Stable Isotope Dilution Liquid Chromatography-Mass Spectrometry Analysis

The metabolism of fatty acids, such as arachidonic acid (AA) and linoleic acid (LA), results in the formation of oxidized bioactive lipids, including numerous stereoisomers1,2. These metabolites can be formed from free or esterified fatty acids. Many of these oxidized metabolites have biological activity and have been implicated in various diseases including cardiovascular and neurodegenerative diseases, asthma, and cancer3-7. Oxidized bioactive lipids can be formed enzymatically or by reactive oxygen species (ROS). Enzymes that metabolize fatty acids include cyclooxygenase (COX), lipoxygenase (LO), and cytochromes P450 (CYPs)1,8. Enzymatic metabolism results in enantioselective formation whereas ROS oxidation results in the racemic formation of products.
While this protocol focuses primarily on the analysis of AA- and some LA-derived bioactive metabolites; it could be easily applied to metabolites of other fatty acids. Bioactive lipids are extracted from cell lysate or media using liquid-liquid (l-l) extraction. At the beginning of the l-l extraction process, stable isotope internal standards are added to account for errors during sample preparation. Stable isotope dilution (SID) also accounts for any differences, such as ion suppression, that metabolites may experience during the mass spectrometry (MS) analysis9. After the extraction, derivatization with an electron capture (EC) reagent, pentafluorylbenzyl bromide (PFB) is employed to increase detection sensitivity10,11. Multiple reaction monitoring (MRM) is used to increase the selectivity of the MS analysis. Before MS analysis, lipids are separated using chiral normal phase high performance liquid chromatography (HPLC). The HPLC conditions are optimized to separate the enantiomers and various stereoisomers of the monitored lipids12. This specific LC-MS method monitors prostaglandins (PGs), isoprostanes (isoPs), hydroxyeicosatetraenoic acids (HETEs), hydroxyoctadecadienoic acids (HODEs), oxoeicosatetraenoic acids (oxoETEs) and oxooctadecadienoic acids (oxoODEs); however, the HPLC and MS parameters can be optimized to include any fatty acid metabolites13.
Most of the currently available bioanalytical methods do not take into account the separate quantification of enantiomers. This is extremely important when trying to deduce whether or not the metabolites were formed enzymatically or by ROS. Additionally, the ratios of the enantiomers may provide evidence for a specific enzymatic pathway of formation. The use of SID allows for accurate quantification of metabolites and accounts for any sample loss during preparation as well as the differences experienced during ionization. Using the PFB electron capture reagent increases the sensitivity of detection by two orders of magnitude over conventional APCI methods. Overall, this method, SID-LC-EC-atmospheric pressure chemical ionization APCI-MRM/MS, is one of the most sensitive, selective, and accurate methods of quantification for bioactive lipids.

This protocol will demonstrate the extraction and analysis of free and esterified bioactive fatty acids from cells. Fatty acids are accurately quantified using stable isotope dilution, chiral liquid chromatography, electron capture atmospheric chemical ionization multiple reaction monitoring mass spectrometry (SID-LC-ECAPCI-MRM/MS).

The metabolism of fatty acids, such as arachidonic acid (AA) and linoleic acid (LA), results in the formation of oxidized bioactive lipids, including ...

Correlative Microscopy for 3D Structural Analysis of Dynamic Interactions

Cryo-electron tomography (cryoET) allows 3D visualization of cellular structures at molecular resolution in a close-to-physiological state1. However, direct visualization of individual viral complexes in their host cellular environment with cryoET is challenging2, due to the infrequent and dynamic nature of viral entry, particularly in the case of HIV-1. While time-lapse live-cell imaging has yielded a great deal of information about many aspects of the life cycle of HIV-13-7, the resolution afforded by live-cell microscopy is limited (~ 200 nm). Our work was aimed at developing a correlation method that permits direct visualization of early events of HIV-1 infection by combining live-cell fluorescent light microscopy, cryo-fluorescent microscopy, and cryoET. In this manner, live-cell and cryo-fluorescent signals can be used to accurately guide the sampling in cryoET. Furthermore, structural information obtained from cryoET can be complemented with the dynamic functional data gained through live-cell imaging of fluorescent labeled target.
In this video article, we provide detailed methods and protocols for structural investigation of HIV-1 and host-cell interactions using 3D correlative high-speed live-cell imaging and high-resolution cryoET structural analysis. HeLa cells infected with HIV-1 particles were characterized first by confocal live-cell microscopy, and the region containing the same viral particle was then analyzed by cryo-electron tomography for 3D structural details. The correlation between two sets of imaging data, optical imaging and electron imaging, was achieved using a home-built cryo-fluorescence light microscopy stage. The approach detailed here will be valuable, not only for study of virus-host cell interactions, but also for broader applications in cell biology, such as cell signaling, membrane receptor trafficking, and many other dynamic cellular processes.

We describe a correlative microscopy method that combines high-speed 3D live-cell fluorescent light microscopy and high-resolution cryo-electron tomography. We demonstrate the capability of the correlative method by imaging dynamic, small HIV-1 particles interacting with host HeLa cells.

Cryo-electron tomography (cryoET) allows 3D visualization of cellular structures at molecular resolution in a close-to-physiological state1. However, ...

Tissue Engineering: Construction of a Multicellular 3D Scaffold for the Delivery of Layered Cell Sheets

Many tissues, such as the adult human hearts, are unable to adequately regenerate after damage.2,3 Strategies in tissue engineering propose innovations to assist the body in recovery and repair. For example, TE approaches may be able to attenuate heart remodeling after myocardial infarction (MI) and possibly increase total heart function to a near normal pre-MI level.4 As with any functional tissue, successful regeneration of cardiac tissue involves the proper delivery of multiple cell types with environmental cues favoring integration and survival of the implanted cell/tissue graft. Engineered tissues should address multiple parameters including: soluble signals, cell-to-cell interactions, and matrix materials evaluated as delivery vehicles, their effects on cell survival, material strength, and facilitation of cell-to-tissue organization. Studies employing the direct injection of graft cells only ignore these essential elements.2,5,6 A tissue design combining these ingredients has yet to be developed. Here, we present an example of integrated designs using layering of patterned cell sheets with two distinct types of biological-derived materials containing the target organ cell type and endothelial cells for enhancing new vessels formation in the &#8220;tissue&#8221;. Although these studies focus on the generation of heart-like tissue, this tissue design can be applied to many organs other than heart with minimal design and material changes, and is meant to be an off-the-shelf product for regenerative therapies. The protocol contains five detailed steps. A temperature sensitive Poly(N-isopropylacrylamide) (pNIPAAM) is used to coat tissue culture dishes. Then, tissue specific cells are cultured on the surface of the coated plates/micropattern surfaces to form cell sheets with strong lateral adhesions. Thirdly, a base matrix is created for the tissue by combining porous matrix with neovascular permissive hydrogels and endothelial cells. Finally, the cell sheets are lifted from the pNIPAAM coated dishes and transferred to the base element, making the complete construct.

For creation of highly organized structures of complex tissue, one must assemble multiple material and cell types into an integrated composite. This combinatorial design incorporates organ-specific layered cell sheets with two distinct biologically-derived materials containing a strong fibrous matrix base, and endothelial cells for enhancing new vessels formation.

Many tissues, such as the adult human hearts, are unable to adequately regenerate after damage.2,3 Strategies in tissue engineering propose innovations to ...

Preparation of 3D Collagen Gels and Microchannels for the Study of 3D Interactions In Vivo

Historically, most cellular processes have been studied in only 2 dimensions. While these studies have been informative about general cell signaling mechanisms, they neglect important cellular cues received from the structural and mechanical properties of the local microenvironment and extracellular matrix (ECM). To understand how cells interact within a physiological ECM, it is important to study them in the context of 3 dimensional assays. Cell migration, cell differentiation, and cell proliferation are only a few processes that have been shown to be impacted by local changes in the mechanical properties of a 3-dimensional ECM. Collagen I, a core fibrillar component of the ECM, is more than a simple structural element of a tissue. Under normal conditions, mechanical cues from the collagen network direct morphogenesis and maintain cellular structures. In diseased microenvironments, such as the tumor microenvironment, the collagen network is often dramatically remodeled, demonstrating altered composition, enhanced deposition and altered fiber organization. In breast cancer, the degree of fiber alignment is important, as an increase in aligned fibers perpendicular to the tumor boundary has been correlated to poorer patient prognosis1. Aligned collagen matrices result in increased dissemination of tumor cells via persistent migration2,3. The following is a simple protocol for embedding cells within a 3-dimensional, fibrillar collagen hydrogel. This protocol is readily adaptable to many platforms, and can reproducibly generate both aligned and random collagen matrices for investigation of cell migration, cell division, and other cellular processes in a tunable, 3-dimensional, physiological microenvironment.

Preparation of 3D Collagen Gels and Microchannels for the Study of 3D Interactions In Vivo

Collagen is a core component of the ECM, and provides essential cues for several cellular processes ranging from migration to differentiation and proliferation. Provided here is a protocol for embedding cells within 3D collagen hydrogels, and a more advanced technique for generating randomized or aligned collagen matrices using PDMS microchannels.

Historically, most cellular processes have been studied in only 2 dimensions.  While these studies have been informative about general cell signaling ...

Bioprintable Alginate/Gelatin Hydrogel 3D In Vitro Model Systems Induce Cell Spheroid Formation

The cellular, biochemical, and biophysical heterogeneity of the native tumor microenvironment is not recapitulated by growing immortalized cancer cell lines using conventional two-dimensional (2D) cell culture. These challenges can be overcome by using bioprinting techniques to build heterogeneous three-dimensional (3D) tumor models whereby different types of cells are embedded. Alginate and gelatin are two of the most common biomaterials employed in bioprinting due to their biocompatibility, biomimicry, and mechanical properties. By combining the two polymers, we achieved a bioprintable composite hydrogel with similarities to the microscopic architecture of a native tumor stroma. We studied the printability of the composite hydrogel via rheology and obtained the optimal printing window. Breast cancer cells and fibroblasts were embedded in the hydrogels and printed to form a 3D model mimicking the in vivo microenvironment. The bioprinted heterogeneous model achieves a high viability for long-term cell culture (&#62; 30 days) and promotes the self-assembly of breast cancer cells into multicellular tumor spheroids (MCTS). We observed the migration and interaction of the cancer-associated fibroblast cells (CAFs) with the MCTS in this model. By using bioprinted cell culture platforms as co-culture systems, it offers a unique tool to study the dependence of tumorigenesis on the stroma composition. This technique features a high-throughput, low cost, and high reproducibility, and it can also provide an alternative model to conventional cell monolayer cultures and animal tumor models to study cancer biology.

Bioprintable Alginate/Gelatin Hydrogel 3D In Vitro Model Systems Induce Cell Spheroid Formation

We developed a heterogeneous breast cancer model consisting of immortalized tumor and fibroblast cells embedded in a bioprintable alginate/gelatin bioink. The model recapitulates the in vivo tumor microenvironment and facilitates the formation of multicellular tumor spheroids, yielding insight into the mechanisms driving tumorigenesis.

The cellular, biochemical, and biophysical heterogeneity of the native tumor microenvironment is not recapitulated by growing immortalized cancer cell ...

Construction of a Multilayered Mesenchymal Stem Cell Sheet with a 3D Dynamic Culture System

Stem cell therapy shows a promising future in regenerating injured organ and tissues, and the cell sheet technique has been developed to improve the low cell retention and poor survival within the target zone. However, during the in vitro construction process, a solution for maintaining stem cell bioactivity and increasing the cell amount within the cell sheet is urgently needed. Here, this protocol presents a method for constructing a multilayered cell sheet with favorable stem cell bioactivity and optimal operability. Decellularized porcine pericardium (DPP) is prepared by phospholipase A2 (PLA2) decellularization method as the cell sheet scaffold, and rat bone marrow mesenchymal stem cells (BMSCs) are isolated and expanded as the seeded cells. The temporary multilayered cell sheet structure is constructed by using RAD16-I peptide hydrogel. Finally, the cell sheet is cultured with a dynamic perfusion system to stabilize the three-dimensional (3D) structure, and the cell sheet could be obtained following a 48-hour culture in vitro. This protocol provides an efficient and feasible method for constructing a multilayered stem cell sheet, and the cell sheet could be developed as a favorable stem cell therapy product in the future.

This article provides an efficient and feasible method for constructing multilayered stem cell sheets with favorable stem cell property.

Stem cell therapy shows a promising future in regenerating injured organ and tissues, and the cell sheet technique has been developed to improve the low ...

Environmental Dynamic Mechanical Analysis to Predict the Softening Behavior of Neural Implants

When using dynamically softening substrates for neural implants, it is important to have a reliable in vitro method to characterize the softening behavior of these materials. In the past, it has not been possible to satisfactorily measure the softening of thin films under conditions mimicking body environment without substantial effort. This publication presents a new and simple method that allows dynamic mechanical analysis (DMA) of polymers in solutions, such as phosphate buffered saline (PBS), at relevant temperatures. The use of environmental DMA allows measurement of the softening effects of polymers due to plasticization in various media and temperatures, which therefore allows a prediction of the materials behavior under in vivo conditions.

To allow reliable predictions of the softening of polymeric substrates for neural implants in an in vivo environment, it is important to have a reliable in vitro method. Here, the use of dynamic mechanical analysis in phosphate buffered saline at body temperature is presented.

When using dynamically softening substrates for neural implants, it is important to have a reliable in vitro method to characterize the softening behavior ...

Ovarian Cancer Detection Using Photoacoustic Flow Cytometry

Many studies suggest that the enumeration of circulating tumor cells (CTCs) may show promise as a prognostic tool for ovarian cancer. Current strategies for the detection of CTCs include flow cytometry, microfluidic devices, and real-time polymerase chain reaction (RT-PCR). Despite recent advances, methods for the detection of early ovarian cancer metastasis still lack the sensitivity and specificity required for clinical translation. Here, a novel method is presented for the detection of ovarian circulating tumor cells by photoacoustic flow cytometry (PAFC) utilizing a custom three dimensional (3D) printed system, including a flow chamber and syringe pump. This method utilizes folic acid-capped copper sulfide nanoparticles (FA-CuS NPs) to target SKOV-3 ovarian cancer cells by PAFC. This work demonstrates the affinity of these contrast agents for ovarian cancer cells. The results show NP characterization, PAFC detection, and NP uptake by fluorescence microscopy, thus demonstrating the potential of this novel system to detect ovarian CTCs at physiologically relevant concentrations.

A protocol is presented to detect circulating ovarian tumor cells utilizing a custom-made photoacoustic flow system and targeted folic acid-capped copper sulfide nanoparticles.

Many studies suggest that the enumeration of circulating tumor cells (CTCs) may show promise as a prognostic tool for ovarian cancer. Current strategies ...

Generating Controlled, Dynamic Chemical Landscapes to Study Microbial Behavior

We demonstrate a method for the generation of controlled, dynamic chemical pulses&#8213;where localized chemoattractant becomes suddenly available at the microscale&#8213;to create micro-environments for microbial chemotaxis experiments. To create chemical pulses, we developed a system to introduce amino acid sources near-instantaneously by photolysis of caged amino acids within a polydimethylsiloxane (PDMS) microfluidic chamber containing a bacterial suspension. We applied this method to the chemotactic bacterium, Vibrio ordalii, which can actively climb these dynamic chemical gradients while being tracked by video microscopy. Amino acids, rendered biologically inert (&#39;caged&#39;) by chemical modification with a photoremovable protecting group, are uniformly present in the suspension but not available for consumption until their sudden release, which occurs at user-defined points in time and space by means of a near-UV-A focused LED beam. The number of molecules released in the pulse can be determined by a calibration relationship between exposure time and uncaging fraction, where the absorption spectrum after photolysis is characterized by using UV-Vis spectroscopy. A nanoporous polycarbonate (PCTE) membrane can be integrated into the microfluidic device to allow the continuous removal by flow of the uncaged compounds and the spent media. A strong, irreversible bond between the PCTE membrane and the PDMS microfluidic structure is achieved by coating the membrane with a solution of 3-aminopropyltriethoxysilane (APTES) followed by plasma activation of the surfaces to be bonded. A computer-controlled system can generate user-defined sequences of pulses at different locations and with different intensities, so as to create resource landscapes with prescribed spatial and temporal variability. In each chemical landscape, the dynamics of bacterial movement at the individual scale and their accumulation at the population level can be obtained, thereby allowing the quantification of chemotactic performance and its effects on bacterial aggregations in ecologically relevant environments.

A protocol for the generation of dynamic chemical landscapes by photolysis within microfluidic and millifluidic setups is presented. This methodology is suitable to study diverse biological processes, including the motile behavior, nutrient uptake, or adaptation to chemicals of microorganisms, both at the single cell and population level.

We demonstrate a method for the generation of controlled, dynamic chemical pulses&#8213;where localized chemoattractant becomes suddenly available at the ...