Development of the model demonstrated here was supported by DOD grant W81XWH-12-1-0163, NIH grants R21 CA179981 and R21 CA196202, and the Sato Metastasis Research Fund.

1. Cell preparation
<ol>
	<li>Release cells from tissue culture dish when 70-90% confluent by following the recommended guidelines for the cell line in use.
	<ol>
		<li>For example, aspirate the growth medium for PC-3 cells, and wash the 10 cm dish of cells with 5 mL of calcium- and magnesium-free phosphate-buffered saline (PBS).</li>
		<li>Aspirate the PBS before adding 1 mL of 0.25% trypsin using manufacturer&#39;s protocol.</li>
		<li>After observing the detachment of the cells under an inverted microscope, add 5 mL of DMEM:F12 medium containing 10% fetal bovine serum to inhibit the trypsin.</li>
	</ol>
	</li>
	<li>Place the cell suspension into a conical tube.</li>
	<li>Determine the cell concentration and total cell number.</li>
	<li>Pellet cells by centrifugation (300 &#215; g for 3 min), aspirate the supernatant, and resuspend cells in serum-free tissue culture medium to 5 &#215; 105 cells/mL. 
	&#8203;NOTE: It is critical that the assay medium contains at least 1.17 mM Ca++ as extracellular Ca++ has been demonstrated to be required for cellular resistance to FSS15.</li>
</ol>
2. Fluid shear stress exposure
<ol>
	<li>Prior to exposing cells to FSS, cut a round-bottom 14 mL polystyrene tube at the 7 mL line. Mix the cell suspension, place 5 mL of the suspension into the cut tube, and collect static control samples. 
	NOTE: The volume needed to collect for the static sample depends on the viability assay used (see step 3).</li>
	<li>Draw the cell suspension into a 5 mL syringe, and attach a 30 G &#189;&#34; needle. Uncap the needle, place the syringe onto a syringe pump, secure the syringe, and set the flow rate to achieve the desired level of FSS. 
	NOTE: Table 1 shows the maximum wall shear stress for different needles and flow rates, as well as the minimum level of FSS depending on cell size (10, 15, and 20 &#181;m). Inspect the needle prior to use to ensure that it is not bent; if uncertain, replace the needle with a new one. Needle integrity can have significant impact on the level of FSS applied.</li>
	<li>Run the syringe pump, and collect the sheared sample in the cut tube at an approximate 45&#176; angle to reduce foaming. Collect a sample depending on the type of viability assay or downstream assay needs.
	<ol>
		<li>Carefully remove the syringe and needle from the syringe pump, and use pliers to remove the needle from the syringe, taking care to not touch the needle. 
		NOTE: Non-beveled needles can be used interchangeably with beveled needles as an additional safety measure.</li>
	</ol>
	</li>
	<li>Draw the sheared suspension back into the syringe, carefully reattach the needle using pliers, and place it back into the syringe pump.</li>
	<li>Repeat steps 2.3 and 2.4 until the cell suspension has been exposed to the desired number of pulses of FSS. 
	&#8203;NOTE: To assess the capacity of cells to resist mechanical destruction from FSS exposure the cell suspension is typically subjected to 10 pulses of FSS. However, it has been demonstrated that cells start to undergo biological adaptations in response to FSS after 2 pulses24.</li>
</ol>
3. Viability measurement
NOTE: Viability can be assessed using enzymatic assays (luciferase, resazurin, and WST-1), counting intact cells, flow cytometry, or by clonogenic assays.
<ol>
	<li>For all measures of viability, collect a sample prior to exposing cells to FSS.
	<ol>
		<li>For enzymatic assays, take duplicate 100 &#181;L aliquots and place them into a 96-well plate.</li>
		<li>For flow cytometry, take one 500 &#181;L aliquot and place it into a 1.5 mL tube.</li>
		<li>For clonogenic assay, collect a 100 &#181;L aliquot.</li>
	</ol>
	</li>
	<li>Enzymatic assay
	<ol>
		<li>Collect 100 &#181;L samples after 1, 2, 4, 6, 8, and 10 pulses of FSS exposure and place them in a 96-well plate.</li>
		<li>Add the desired substrate, and follow the protocol for the assay used:
		<ol>
			<li>For resazurin, add 20 &#181;L of a 0.15 mg/mL solution to each well. Add 20 &#181;L of 0.15 mg/mL resazurin solution to wells containing 100 &#181;L of medium alone. Incubate for 2 h in a 37 &#176;C tissue culture incubator. Measure the absorbance using a plate reader capable of reading fluorescence (579 excitation/ 584 emission).</li>
			<li>For luciferase-expressing cells, add 100 &#181;L of 15 mg/mL D-luciferin to 5 mL of medium. Add 100 &#181;L of that solution to each well containing cells. Wait for 5 min, and then read the plate using a reader compatible with luminescence.</li>
			<li>For WST-1, add 10 &#181;L of WST-1 to each well, including wells containing medium only. Incubate for 4 h, and then read the absorbance between 420 and 480 nm using a plate reader.</li>
		</ol>
		</li>
		<li>Compare the averaged signal from each of the FSS-exposed samples to the averaged static control sample to obtain the percentage of viable cells.</li>
	</ol>
	</li>
	<li>Flow cytometry24
	<ol>
		<li>Collect 500 &#181;L samples and place them into 1.5 mL centrifuge tubes after 1, 2, 5, and 10 pulses of FSS.</li>
		<li>Centrifuge samples (500 &#215; g for 3 min), and discard the supernatants.</li>
		<li>Resuspend the pellets with 1 mL of calcium- and magnesium-free PBS, and centrifuge the samples (300 &#215; g for 3 min).</li>
		<li>Suspend the pellets with 500 &#181;L of fluorescence-activated cell sorting (FACS) buffer (PBS with 0.5% bovine serum albumin and 0.1% sodium&#160;azide) with counting beads and membrane-impermeable or viability dyes such as propidium iodide (1.75 &#181;g/mL).</li>
		<li>Determine the viability by comparing the ratio of viable cells, normalized to counting beads, in sheared samples to that of the static sample.</li>
	</ol>
	</li>
	<li>Clonogenic assay
	<ol>
		<li>Take 100 &#181;L of the static sample, and add 900 &#181;L of growth medium to make a 1:10 dilution.</li>
		<li>Take 100 &#181;L of the 1:10 diluted sample, and add 900 &#181;L of growth medium to make a final 1:100 dilution.</li>
		<li>Add 100 &#181;L of the 1:100 dilution sample into each of 3 wells of a 6-well dish containing 2 mL of growth medium.</li>
		<li>Repeat steps 3.4.1-3.4.3 with samples that have been subjected to 10 pulses of FSS.</li>
		<li>Let the cells grow for 7-10 days without changing the medium, and check for colony formation. Once colonies of &#8805;50 cells have formed, aspirate the growth medium, rinse each well with 1 mL of PBS, aspirate the PBS, and fix for 5 min using 1 mL of ice-cold 70% ethanol (EtOH). Importantly, fix both sheared and static samples at the same time</li>
		<li>After fixing the samples, aspirate the EtOH, and add 1 to 2 mL of crystal violet solution (0.1% crystal violet in 90% H2O, 10% EtOH) for 5 min.</li>
		<li>Rinse with an excess of water, and let the plate dry</li>
		<li>Count the colonies (clusters of &#8805;50 cells) for both the static and sheared samples. Compare the ratio of the average number of colonies from the sheared sample to the average number of colonies from the static sample to determine viability.</li>
	</ol>
	</li>
</ol>

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MDH is a co-founder, President and shareholder of SynderBio, Inc. DLM is a consultant for SynderBio, Inc.

This paper demonstrates the application of FSS to cancer cells in suspension using a syringe and needle. Using this model, cancer cells have been shown to be more resistant to brief pulses of high-level FSS relative to non-transformed epithelial cells15,22,24. Furthermore, exposure to FSS using this model results in a rapid increase in cell stiffness, activation of RhoA, and increased cortical F-actin and myosin II-based contrac...

Metastasis, or the spread of cancer beyond the initial tumor site, is a major factor underlying cancer mortality1. During metastasis, cancer cells utilize the circulatory system as a highway to disseminate to distant sites throughout the body2,3. While en route to these sites, circulating tumor cells (CTCs) exist within a dynamic fluid microenvironment unlike that of their original primary tumor3,4,5. It has been proposed that this fluid microenvironment is one of many barriers to metastasis4. There is wide agreement in the concept of metastatic inefficiency, i.e., that most CTCs entering the circulation either perish or do not form productive metastatic colonies6,7,8. However, why metastasis is inefficient from the perspective of an individual CTC is less certain and remains an active area of investigation. CTCs are detached from extracellular matrix, deprived of soluble growth and survival factors that may be present in the primary tumor, and exposed to the immune system and hemodynamic forces in a much different manner than in the primary tumor4. Each of these factors may contribute to the poor survival of CTCs, but their relative contributions are unclear. This paper addresses the question of how hemodynamic forces affect CTCs.
Studying the effects of hemodynamic forces on CTCs is quite challenging. Currently, there are no engineered in vitro systems that can replicate the entire spatiotemporal dynamics (heart to capillaries) and rheological properties of the human vascular system. Moreover, how CTCs experience the circulatory system is not entirely clear. Experimental evidence indicates that most cancer cells do not circulate continuously like blood cells. Rather, due to their relatively large size (10-20 &#181;m in diameter), most CTCs become entrapped in capillary beds (6-8 &#181;m in diameter) for variable lengths of time (s to days) where they may die, extravasate, or be displaced to the next capillary bed8,9,10,11. However, there is some evidence that CTC size may be more heterogeneous in vivo, and that smaller CTCs are detectable12. Therefore, based on distance and blood flow velocity, CTCs may only circulate freely for a matter of seconds between these periods of entrapment, although a quantitative description of this behavior is lacking13.
Furthermore, depending on where CTCs enter the circulation, they may pass through multiple capillary beds in the lung and other peripheral sites and through both the right and left heart prior to reaching their final destination. Along the way, CTCs are exposed to various hemodynamic stresses including fluid shear stress (FSS), compressive forces during their entrapment in the microcirculation, and potentially, traction forces under circumstances where they might exhibit leukocyte-like rolling along blood vessel walls14. Thus, both the ability to model the circulation and the understanding of the CTC behavior to be modeled is limited. Because of this uncertainty, any findings from in vitro model systems should be validated in an experimental vertebrate organism and ultimately, in cancer patients.
With the aforementioned caveats, this paper demonstrates a relatively simple model to apply FSS to cells in suspension to probe the effects of FSS on CTCs first described in 201215. FSS results from friction of blood flow against the vessel wall, which produces a parabolic velocity gradient under conditions of laminar flow in larger vessels. Cells experience higher levels of FSS near vessel walls and lower levels near the center of the blood vessel. Fluid viscosity, flow rate, and dimensions of the conduit through which the flow occurs influence FSS, as described by the Hagen-Poiseuille equation. This applies to blood flows behaving as Newtonian fluids, but does not hold for the microcirculation. Physiological FSS ranges over several orders of magnitude with the lowest levels in the lymphatics (&#60;1 dyn/cm2) and the highest at regions around heart valves and atherosclerotic plaques (&#62;500 dyn/cm2)5. Mean wall shear stress in arteries is 10-70 dyn/cm2 and 1-6 dyn/cm2 in veins16,17.
In the heart, cells may be exposed to turbulent flows around valve leaflets where very high-level, but very short-duration FSS may be experienced18,19. Although the bioprocessing field has long studied the effects of FSS on mammalian cells in suspension, this information may be of limited value for understanding the effects of FSS on CTCs as it generally focuses on much lower levels of FSS applied over a long duration20. As described below, using a syringe and needle, one can apply relatively high (tens to thousands dyn/cm2) FSS for a relatively short (milliseconds) duration to a cell suspension. Since the initial description of this model15, others have employed it to study the effects of FSS on cancer cells21,22,23. Multiple &#34;pulses&#34; of FSS can be applied to cell suspensions in a short period of time to facilitate downstream experimental analyses. For example, this model can be used to measure the ability of cells to resist mechanical destruction by FSS by measuring cell viability as a function of the number of pulses applied. Alternatively, the effects of FSS exposure on the biology of cancer cells can be explored by collecting cells for a variety of downstream analyses. Importantly, part of the cell suspension is reserved as a static control to compare the effects of FSS from those that might be associated with cell detachment and time held in suspension.

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			<td>Gibco</td>
			<td>25200-056</td>
			<td></td>
		</tr>
		<tr>
			<td>14 mL round bottom tubes</td>
			<td>Falcon - Corning</td>
			<td>352059</td>
			<td></td>
		</tr>
		<tr>
			<td>30 G 1/2&#34; Needle</td>
			<td>BD</td>
			<td>305106</td>
			<td></td>
		</tr>
		<tr>
			<td>5 mL syringe</td>
			<td>BD</td>
			<td>309646</td>
			<td></td>
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			<td>96-well black bottom plate</td>
			<td>Costar - Corning</td>
			<td>3915</td>
			<td></td>
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		<tr>
			<td>Bioluminescence detector</td>
			<td>AMI</td>
			<td>AMI HTX</td>
			<td></td>
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		<tr>
			<td>BSA, Fraction V</td>
			<td>Sigma</td>
			<td>10735086001</td>
			<td></td>
		</tr>
		<tr>
			<td>Cell Titer Blue</td>
			<td>Promega</td>
			<td>G8081</td>
			<td></td>
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			<td>crystal violet</td>
			<td>Sigma</td>
			<td>C0775</td>
			<td></td>
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		<tr>
			<td>D-luciferin</td>
			<td>GoldBio</td>
			<td>D-LUCK</td>
			<td></td>
		</tr>
		<tr>
			<td>DMEM</td>
			<td>Gibco</td>
			<td>11965-092</td>
			<td></td>
		</tr>
		<tr>
			<td>FBS</td>
			<td>Atlanta Biologicals</td>
			<td>S11150</td>
			<td></td>
		</tr>
		<tr>
			<td>PBS</td>
			<td>Gibco</td>
			<td>10010023</td>
			<td></td>
		</tr>
		<tr>
			<td>Plate Reader</td>
			<td>BioTek</td>
			<td>Synergy HT</td>
			<td></td>
		</tr>
		<tr>
			<td>Sodium Azide (NaN3)</td>
			<td>Sigma</td>
			<td>S2002</td>
			<td></td>
		</tr>
		<tr>
			<td>Syringe Pump</td>
			<td>Harvard Apparatus</td>
			<td>70-3005</td>
			<td></td>
		</tr>
	</tbody>
</table>

modeling the effects of hemodynamic stress on circulating tumor cells using a syringe and needle

During metastasis, cancer cells from solid tissues, including epithelia, gain access to the lymphatic and hematogenous circulation where they are exposed to mechanical stress due to hemodynamic flow. One of these stresses that circulating tumor cells (CTCs) experience is fluid shear stress (FSS). While cancer cells may experience low levels of FSS within the tumor due to interstitial flow, CTCs are exposed, without extracellular matrix attachment, to much greater levels of FSS. Physiologically, FSS ranges over 3-4 orders of magnitude, with low levels present in lymphatics (&#60;1 dyne/cm2) and the highest levels present briefly as cells pass through the heart and around heart valves (&#62;500 dynes/cm2). There are a few in vitro models designed to model different ranges of physiological shear stress over various time frames. This paper describes a model to investigate the consequences of brief (millisecond) pulses of high-level FSS on cancer cell biology using a simple syringe and needle system.

Elevated resistance to FSS-induced mechanical destruction has been previously shown to be a conserved phenotype across multiple cancer cell lines and cancer cells freshly isolated from tumors relative to non-transformed epithelial cell comparators15,24. Here, additional cancer cell lines from a variety of tissue origins (Table 2) were tested to demonstrate that the majority of these cells display viability &#8805; 20% after 10 pulses of FSS at 25...

Watch this Scientific Journal Video about Modeling the Effects of Hemodynamic Stress on Circulating Tumor Cells using a Syringe and Needle at JoVE.com

Modeling the Effects of Hemodynamic Stress on Circulating Tumor Cells using a Syringe and Needle

Here we demonstrate a method to apply fluid shear stress to cancer cells in suspension to model the effects of hemodynamic stress on circulating tumor cells.

During metastasis, cancer cells from solid tissues, including epithelia, gain access to the lymphatic and hematogenous circulation where they are exposed ...

modeling-effects-hemodynamic-stress-on-circulating-tumor-cells-using

Research

JoVE Journal

Cancer Research

2.5K Views.  University of Iowa. Here we demonstrate a method to apply fluid shear stress to cancer cells in suspension to model the effects of hemodynamic stress on circulating tumor cells. 

Video: Modeling the Effects of Hemodynamic Stress on Circulating Tumor Cells using a Syringe and Needle

Capture and Release of Viable Circulating Tumor Cells from Blood

We demonstrate a method for size based capture of viable circulating tumor cell (CTC) from whole blood, along with the release of these cells from chip for downstream analysis and/or culture. The strategy employs the use of a novel Parylene C membrane slot pore microfilter to capture CTC and a coating of poly (N-iso-propylacrylamide) (PIPAAm) for thermoresponsive viable release of the captured CTC. The capture of live cells is enabled by leveraging the design of a slot pore geometry with specific dimensions to reduce the shear stress typically associated with the filtration process. While the microfilter exhibits a high capture efficiency, the release of these cells is non-trivial. Typically, only a small percentage of cells are released when techniques such as reverse flow or cell scraping are used. The strong adhesion of these epithelial cancer cells to the Parylene C membrane is attributable to non-specific electrostatic interaction. To counteract this effect, we employed the use of PIPAAm coating and exploited its thermal responsive interfacial properties to release the cells from the filter. Blood is first filtered at room temperature. Below 32 &#176;C, PIPAAm is hydrophilic. Thereafter, the filter is placed in either culture media or a buffer maintained at 37 &#176;C, which results in the PIPAAm turning hydrophobic, and subsequently releasing the electrostatically bound cells.

A protocol to utilize a poly(N-iso-propylacrylamide) (PIPAAm) coated microfilter for effective capture and thermoresponsive release of viable circulating tumor cells (CTC) is presented. This method allows capture of CTC from patients&#39; blood and subsequent release of viable CTC for downstream off-chip culture, analyses and characterization.

We demonstrate a method for size based capture of viable circulating tumor cell (CTC) from whole blood, along with the release of these cells from chip ...

The Drosophila Imaginal Disc Tumor Model: Visualization and Quantification of Gene Expression and Tumor Invasiveness Using Genetic Mosaics

Drosophila melanogaster has emerged as a powerful experimental system for functional and mechanistic studies of tumor development and progression in the context of a whole organism. Sophisticated techniques to generate genetic mosaics facilitate induction of visually marked, genetically defined clones surrounded by normal tissue. The clones can be analyzed through diverse molecular, cellular and omics approaches. This study describes how to generate fluorescently labeled clonal tumors of varying malignancy in the eye/antennal imaginal discs (EAD) of Drosophila larvae using the Mosaic Analysis with a Repressible Cell Marker (MARCM) technique. It describes procedures how to recover the mosaic EAD and brain from the larvae and how to process them for simultaneous imaging of fluorescent transgenic reporters and antibody staining. To facilitate molecular characterization of the mosaic tissue, we describe a protocol for isolation of total RNA from the EAD. The dissection procedure is suitable to recover EAD and brains from any larval stage. The fixation and staining protocol for imaginal discs works with a number of transgenic reporters and antibodies that recognize Drosophila proteins. The protocol for RNA isolation can be applied to various larval organs, whole larvae, and adult flies. Total RNA can be used for profiling of gene expression changes using candidate or genome-wide approaches. Finally, we detail a method for quantifying invasiveness of the clonal tumors. Although this method has limited use, its underlying concept is broadly applicable to other quantitative studies where cognitive bias must be avoided.

The Drosophila Imaginal Disc Tumor Model: Visualization and Quantification of Gene Expression and Tumor Invasiveness Using Genetic Mosaics

This protocol demonstrates how to generate fluorescently marked, genetically defined clonal tumors in the Drosophila eye/antennal imaginal discs (EAD). It describes how to dissect the EAD and brain from the third instar larvae and how to process them to visualize and quantify gene expression changes and tumor invasiveness.

Drosophila melanogaster has emerged as a powerful experimental system for functional and mechanistic studies of tumor development and progression in the ...

Isolation of Circulating Tumor Cells in an Orthotopic Mouse Model of Colorectal Cancer

Despite the advantages of easy applicability and cost-effectiveness, subcutaneous mouse models have severe limitations and do not accurately simulate tumor biology and tumor cell dissemination. Orthotopic mouse models have been introduced to overcome these limitations; however, such models are technically demanding, especially in hollow organs such as the large bowel. In order to produce uniform tumors which reliably grow and metastasize, standardized techniques of tumor cell preparation and injection are critical. 
We have developed an orthotopic mouse model of colorectal cancer (CRC) which develops highly uniform tumors and can be used for tumor biology studies as well as therapeutic trials. Tumor cells from either primary tumors, 2-dimensional (2D) cell lines or 3-dimensional (3D) organoids are injected into the cecum and, depending on the metastatic potential of the injected tumor cells, form highly metastatic tumors. In addition, CTCs can be found regularly. We here describe the technique of tumor cell preparation from both 2D cell lines and 3D organoids as well as primary tumor tissue, the surgical and injection techniques as well as the isolation of CTCs from the tumor-bearing mice, and present tips for troubleshooting.

We describe the establishment of orthotopic colorectal tumors via injection of tumor cells or organoids into the cecum of mice and the subsequent isolation of circulating tumor cells (CTCs) from this model.

Despite the advantages of easy applicability and cost-effectiveness, subcutaneous mouse models have severe limitations and do not accurately simulate ...

The Establishment of a Lung Colonization Assay for Circulating Tumor Cell Visualization in Lung Tissues

Metastasis is the major cause of cancer death. The role of circulating tumor cells (CTCs) in promoting cancer metastasis, in which lung colonization by CTCs critically contributes to early lung metastatic processes, has been vigorously investigated. As such, animal models are the only approach that captures the full systemic process of metastasis. Given that problems occur in previous experimental designs for examining the contributions of CTCs to blood vessel extravasation, we established an in vivo lung colonization assay in which a long-term-fluorescence cell-tracer, carboxyfluorescein succinimidyl ester (CFSE), was used to label suspended tumor cells and lung perfusion was performed to clear non-specifically trapped CTCs prior to lung removal, confocal imaging, and quantification. Polymeric fibronectin (polyFN) assembled on CTC surfaces has been found to mediate lung colonization in the final establishment of metastatic tumor tissues. Here, to specifically test the requirement of polyFN assembly on CTCs for lung colonization and extravasation, we performed short term lung colonization assays in which suspended Lewis lung carcinoma cells (LLCs) stably expressing FN-shRNA (shFN) or scramble-shRNA (shScr) and pre-labeled with 20 &#956;M of CFSE were intravenously inoculated into C57BL/6 mice. We successfully demonstrated that the abilities of shFN LLC cells to colonize the mouse lungs were significantly diminished in comparison to shScr LLC cells. Therefore, this short-term methodology may be widely applied to specifically demonstrate the ability of CTCs within the circulation to colonize the lungs.

An animal model is needed to decipher the role of circulating tumor cells (CTCs) in promoting lung colonization during cancer metastasis. Here, we established and successfully performed an in vivo assay to specifically test the requirement of polymeric fibronectin (polyFN) assembly on CTCs for lung colonization.

Metastasis is the major cause of cancer death. The role of circulating tumor cells (CTCs) in promoting cancer metastasis, in which lung colonization by ...

Characterization of Tumor Cells Using a Medical Wire for Capturing Circulating Tumor Cells: A 3D Approach Based on Immunofluorescence and DNA FISH

Circulating tumor cells (CTCs) are associated with poor survival in metastatic cancer. Their identification, phenotyping, and genotyping could lead to a better understanding of tumor heterogeneity and thus facilitate the selection of patients for personalized treatment. However, this is hampered because of the rarity of CTCs. We present an innovative approach for sampling a high volume of the patient blood and obtaining information about presence, phenotype, and gene translocation of CTCs. The method combines immunofluorescence staining and DNA fluorescent-in-situ-hybridization (DNA FISH) and is based on a functionalized medical wire. This wire is an innovative device that permits the in vivo isolation of CTCs from a large volume of peripheral blood. The blood volume screened by a 30-min administration of the wire is approximately 1.5-3 L. To demonstrate the feasibility of this approach, epithelial cell adhesion molecule (EpCAM) expression and the chromosomal translocation of the ALK gene were determined in non-small-cell lung cancer (NSCLC) cell lines captured by the functionalized wire and stained with an immuno-DNA FISH approach. Our main challenge was to perform the assay on a 3D structure, the functionalized wire, and to determine immuno-phenotype and FISH signals on this support using a conventional fluorescence microscope. The results obtained indicate that catching CTCs and analyzing their phenotype and chromosomal rearrangement could potentially represent a new companion diagnostic approach and provide an innovative strategy for improving personalized cancer treatments.

We present a new approach to characterize tumor cells. We combined immunofluorescence with DNA fluorescent-in-situ-hybridization to evaluate cells captured by a functionalized medical wire capable of in vivo enriching CTCs directly from patient blood.

Circulating tumor cells (CTCs) are associated with poor survival in metastatic cancer. Their identification, phenotyping, and genotyping could lead to a ...

Imaging Approaches to Assessments of Toxicological Oxidative Stress Using Genetically-encoded Fluorogenic Sensors

While oxidative stress is a commonly cited toxicological mechanism, conventional methods to study it suffer from a number of shortcomings, including destruction of the sample, introduction of potential artifacts, and a lack of specificity for the reactive species involved. Thus, there is a current need in the field of toxicology for non-destructive, sensitive, and specific methods that can be used to observe and quantify intracellular redox perturbations, more commonly referred to as oxidative stress. Here, we present a method for the use of two genetically-encoded fluorogenic sensors, roGFP2 and HyPer, to be used in live-cell imaging studies to observe xenobiotic-induced oxidative responses. roGFP2 equilibrates with the glutathione redox potential (EGSH), while HyPer directly detects hydrogen peroxide (H2O2). Both sensors can be expressed into various cell types via transfection or transduction, and can be targeted to specific cellular compartments. Most importantly, live-cell microscopy using these sensors offers high spatial and temporal resolution that is not possible using conventional methods. Changes in the fluorescence intensity monitored at 510 nm serves as the readout for both genetically-encoded fluorogenic sensors when sequentially excited by 404 nm and 488 nm light. This property makes both sensors ratiometric, eliminating common microscopy artifacts and correcting for differences in sensor expression between cells. This methodology can be applied across a variety of fluorometric platforms capable of exciting and collecting emissions at the prescribed wavelengths, making it suitable for use with confocal imaging systems, conventional wide-field microscopy, and plate readers. Both genetically-encoded fluorogenic sensors have been used in a variety of cell types and toxicological studies to monitor cellular EGSH and H2O2 generation in real-time. Outlined here is a standardized method that is widely adaptable across cell types and fluorometric platforms for the application of roGFP2 and HyPer in live-cell toxicological assessments of oxidative stress.

This manuscript describes the use of genetically-encoded fluorogenic reporters in an application of live-cell imaging for the examination of xenobiotic-induced oxidative stress. This experimental approach offers unparalleled spatiotemporal resolution, sensitivity, and specificity while avoiding many of the shortcomings of conventional methods used for the detection of toxicological oxidative stress.

While oxidative stress is a commonly cited toxicological mechanism, conventional methods to study it suffer from a number of shortcomings, including ...

A Method to Define the Effects of Environmental Enrichment on Colon Microbiome Biodiversity in a Mouse Colon Tumor Model

Several recent studies have illustrated the beneficial effects of living in an enriched environment on improving human disease. In mice, environmental enrichment (EE) reduces tumorigenesis by activating the mouse immune system, or affects tumor bearing animal survival by stimulating the wound repair response, including improved microbiome diversity, in the tumor microenvironment. Provided here is a detailed procedure to assess the effects of environmental enrichment on the biodiversity of the microbiome in a mouse colon tumor model. Precautions regarding animal breeding and considerations for animal genotype and mouse colony integration are described, all of which ultimately affect microbial biodiversity. Heeding these precautions may allow more uniform microbiome transmission, and consequently will alleviate non-treatment dependent effects that can confound study findings. Further, in this procedure, microbiota changes are characterized using 16S rDNA sequencing of DNA isolated from stool collected from the distal colon following long-term environmental enrichment. Gut microbiota imbalance is associated with the pathogenesis of inflammatory bowel disease and colon cancer, but also of obesity and diabetes among others. Importantly, this protocol for EE and microbiome analysis can be utilized to study the role of microbiome pathogenesis across a variety of diseases where robust mouse models exist that can recapitulate human disease.

Environmental Enrichment (EE) is an animal housing environment that is used to reveal mechanisms that underlie the connections between lifestyle, stress, and disease. This protocol describes a procedure that uses a mouse model of colon tumorigenesis and EE to specifically define alterations in microbiota biodiversity that may impact animal mortality.

Several recent studies have illustrated the beneficial effects of living in an enriched environment on improving human disease. In mice, environmental ...

Modeling Osteosarcoma Using Li-Fraumeni Syndrome Patient-derived Induced Pluripotent Stem Cells

Li-Fraumeni syndrome (LFS) is an autosomal dominant hereditary cancer disorder. Patients with LFS are predisposed to a various type of tumors, including osteosarcoma--one of the most frequent primary non-hematologic malignancies in the childhood and adolescence. Therefore, LFS provides an ideal model to study this malignancy. Taking advantage of iPSC methodologies, LFS-associated osteosarcoma can be successfully modeled by differentiating LFS patient iPSCs to mesenchymal stem cells (MSCs), and then to osteoblasts--the cells of origin for osteosarcomas. These LFS osteoblasts recapitulate oncogenic properties of osteosarcoma, providing an attractive model system for delineating the pathogenesis of osteosarcoma. This manuscript demonstrates a protocol for the generation of iPSCs from LFS patient fibroblasts, differentiation of iPSCs to MSCs, differentiation of MSCs to osteoblasts, and in vivo tumorigenesis using LFS osteoblasts. This iPSC disease model can be extended to identify potential biomarkers or therapeutic targets for LFS-associated osteosarcoma.

Here, we present a protocol for the generation of induced pluripotent Stem Cells (iPSCs) from Li-Fraumeni Syndrome (LFS) patient derived fibroblasts, differentiation of iPSCs via mesenchymal stem cells (MSCs) to osteoblasts, and modeling in vivo tumorigenesis using LFS patient-derived osteoblasts.

Li-Fraumeni syndrome (LFS) is an autosomal dominant hereditary cancer disorder. Patients with LFS are predisposed to a various type of tumors, including ...

Digital Polymerase Chain Reaction Assay for the Genetic Variation in a Sporadic Familial Adenomatous Polyposis Patient Using the Chip-in-a-tube Format

The quantitative analysis of human genetic variation is crucial for understanding the molecular characteristics of serious medical conditions, such as tumors. Because digital polymerase chain reactions (PCR) enable the precise quantification of DNA copy number variants, they are becoming an essential tool for detecting rare genetic variations, such as drug-resistant mutations. It is expected that molecular diagnoses using digital PCR (dPCR) will be available in clinical practice in the near future; thus, how to efficiently conduct dPCR with human genetic material is a hot topic. Here, we introduce a method to detect Adenomatous polyposis coli (APC) somatic mosaicism using dPCR with the chip-in-a-tube format, which allows eight dPCR reactions to be simultaneously conducted. Care should be taken when filling and sealing the reaction mixture on the chips. This article demonstrates how to avoid the over- and underestimation of positive partitions. Furthermore, we present a simple procedure for collecting the dPCR product from the partitions on the chips, which can then be used to confirm the specific amplification. We hope that this methods report will help promote the dPCR with the chip-in-a-tube method in genetic research.

Digital polymerase chain reaction (PCR) is a useful tool for the high-sensitivity detection of single nucleotide variants and DNA copy number variants. Here, we demonstrate key considerations for measuring rare variants in the human genome using digital PCR with the chip-in-a-tube format.

The quantitative analysis of human genetic variation is crucial for understanding the molecular characteristics of serious medical conditions, such as ...

Micromanipulation of Circulating Tumor Cells for Downstream Molecular Analysis and Metastatic Potential Assessment

Blood-borne metastasis accounts for&#160;most cancer-related deaths and involves circulating tumor cells (CTCs) that are successful in establishing new tumors at distant sites. CTCs are found in the bloodstream of patients as single cells (single CTCs) or as multicellular aggregates (CTC clusters and CTC-white blood cell clusters), with the latter displaying a higher metastatic ability. Beyond enumeration, phenotypic and molecular analysis is extraordinarily important to dissect CTC biology and to identify actionable vulnerabilities. Here, we provide a detailed description of a workflow that includes CTC immunostaining and micromanipulation, ex vivo culture to assess&#160;proliferative and survival capabilities of individual cells, and in vivo metastasis-formation assays. Additionally, we provide a protocol to achieve the dissociation of CTC clusters into individual cells and the investigation of intra-cluster heterogeneity. With these approaches, for instance, we precisely quantify survival and proliferative potential of single CTCs and individual cells within CTC clusters, leading us to the observation that cells within clusters display better survival and proliferation in ex vivo cultures compared to single CTCs. Overall, our workflow offers a platform to dissect the characteristics of CTCs at the single cell level, aiming towards the identification of metastasis-relevant pathways and a better understanding of CTC biology.

Here, we present an integrated workflow to identify phenotypic and molecular features that characterize circulating tumor cells (CTCs). We combine live immunostaining and robotic micromanipulation of single and clustered CTCs with single cell-based techniques for downstream analysis and assessment of metastasis-seeding ability.

Blood-borne metastasis accounts for&#160;most cancer-related deaths and involves circulating tumor cells (CTCs) that are successful in establishing new ...