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 contractility24,27. Rapid mechano-adaptation (the ability of CTCs to become more or less stiff depending on the circumstances) may prevent the mechanical destruction of CTCs and facilitate other aspects of metastatic colonization24,28. Indeed, findings made using this in vitro model have been confirmed using experimental CTCs in animal models24. This rapid mechano-adaptation likely explains the bi-phasic loss of cell viability typically observed in this model (Figure 1), i.e., FSS-na&#239;ve cells are more susceptible to destruction than cells that have been exposed to even a single pulse of FSS. Taken together, this indicates that FSS induces rapid cell stiffening in cancer cells that protects them from subsequent pulses of FSS.
Although the RhoA-actomyosin axis is an important driver of FSS resistance15,21,24, there are likely other mechanisms involved29. Further evidence that cell stiffness is a key determinant of FSS resistance is that disruption of lamin A, which controls the structural integrity of the nucleus-the stiffest component of the cell, reduces FSS resistance in cancer cells using this model22. We are using this model to probe the mechanisms of FSS resistance in cancer cells further. Here, this model has been used to measure the capacity of various cancer cell lines to resist mechanical destruction by exposing cells to brief pulses of high levels of FSS. Although this is a relatively inexpensive, simple model to develop in the laboratory, with the most expensive element being the syringe pump, care must be taken to follow the protocol faithfully to obtain reproducible results. Multiple pulses of FSS can be applied to cells in a very short time, &#60;10min. The total elapsed time for the experiment depends on the suspension volume, flow rate, pulse number, and the dexterity of the user transferring the suspension between pulses. With experience, a 5 mL suspension exposed to FSS with a 30 G needle for 10 pulses@250 &#181;L/s can be processed in ~10 min. For most cell lines, there is minimal loss of viability due to being held in suspension for this length of time.
Because the exposure to FSS occurs relatively quickly, FSS is typically applied to cell suspensions in serum-free medium to reduce foaming of the samples. The difference in viscosity between 0-10% fetal bovine serum is negligible in this assay. However, it is critical to ensure physiologic levels of calcium in the medium in which the cells are sheared. Moreover, with regard to the methods for cell dissociation prior to FSS exposure, no difference in FSS resistance was detected in PC-3 cell suspensions prepared by trypsinization or treatment with non-enzymatic dissociation agents15. Cell concentration can be greater or less than 5 &#215; 105 cells/mL depending on downstream application needs. The response of PC-3 prostate cancer cells is similar in a range from 5 &#215; 104 to 5 &#215; 105,15. However, the effects of cell concentration on viability after FSS exposure should be empirically determined.
For most applications envisioned with cultured cells, cell density should not significantly affect viscosity and therefore, the amount of FSS applied. Variables, such as the time for which the cells are held in suspension prior to FSS exposure, should be held constant across experimental replicates. As mentioned above, needle integrity is also critical. Lot variations have been noted in needles with respect to this assay over time. Hypodermic needles were designed for clinical use, not for the flow rates employed here. On rare occasions, the hub of the needle can be partially occluded, which during subsequent pulses, occludes the passage of suspension through the needle and ultimately, backflow around the syringe plunger. Further, it is very important to understand that dead/dying cells are exceptionally sensitive to FSS, as shown previously24. Therefore, if a particular cell line has a high level of dying cells, either as a routine characteristic or experimental manipulations (e.g., drug treatments), this will result in a very steep loss of cell viability that might not be completely normalized by comparison to the static control.
The application of FSS can be paired with other assays, such as immunofluorescence, pulldown assays, and western blotting, to study the effect of FSS on cancer cell biology 24. In principle, this model might also be used to explore the effects of high-level, short-duration FSS on other cell types including blood cells. Normal red blood cells and leukocytes are much more resistant to FSS applied in this way than even cancer cells, which stands to reason physiologically15. In fact, the level of FSS applied, using a 30 G &#189;&#34; needle at a flow rate of 250 mL/s, brackets the range required for the disruption of the red cell membrane (based on millisecond application of force)30,31. One limitation of this model, or any that involve passing fluid through a conduit, is that the precise level of FSS that cells experience within the range from the maximum wall shear stress and the minimum at the center of the conduit is not known. Thus, at each pulse, all the cells do not experience the same level of FSS, and over repeated pulses, individual cells would be expected to experience different levels of FSS at each pulse within the range specified.
However, hydrodynamic focusing under the conditions employed in this model results in cells being directed toward the center of the flow, away from the wall, and thus toward lower FSS exposure32. Other models, such as cone and plate viscometers or Couette chambers, are better suited for the application of FSS at constant levels to a cell suspension. As mentioned above, it remains challenging to model FSS exposure of CTCs in vitro. This model is best suited to test the effects of high, but brief, exposure to FSS as might happen traversing the heart. Flow through arteries and veins results in longer exposure to lower levels of FSS. However, as mentioned, how long CTCs remain in continuous flow in the circulation is unclear, and most experimental evidence to date is consistent with short periods (seconds) of free flow punctuated by longer periods of entrapment in the microcirculation.
Models that expose cancer cells in suspension to lower levels of FSS (0.5-60 dyn/cm2) for longer durations (minutes to days) include cone and plate viscometers, Couette chambers, continuous flow loops, syringe with a tube extension, and microfluidic devices33,34,35,36,37. These have also been used to gain insights into how FSS might affect CTCs and have led to finding that exposure to FSS increases oxidative stress, cell proliferation and invasion, and stem cell-like characteristics in various cancer cell lines. It will be interesting to compare results derived from those models with the one described here. For example, using a continuous flow loop model, Xin et al. found that the ROCK-actomyosin axis promoted a loss of cell viability in cancer cell lines exposed to FSS (20 dyn/cm2) for 2-12h in stark contrast to the data described above38. Thus, biological context is very likely to matter for all of these in vitro models, reinforcing the need to translate findings about CTCs into in vivo models and ultimately, cancer patients.

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.

<table><tbody><tr><td>0.25% Trypsin</td><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&quot; 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></tr><tr><td>96-well black bottom plate</td><td>Costar - Corning</td><td>3915</td><td></td></tr><tr><td>Bioluminescence detector</td><td>AMI</td><td>AMI HTX</td><td></td></tr><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></tr><tr><td>crystal violet</td><td>Sigma</td><td>C0775</td><td></td></tr><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 (NaN&lt;sub&gt;3&lt;/sub&gt;)</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 250 &#181;L/s. The one exception is MiaPaCa2 cells, which were relatively sensitive to mechanical destruction from FSS (viability &#8804; 10%). To adequately describe the FSS resistance profile of a cell line, n &#8805; 3 biological replicates are recommended.
By way of comparison, all of the non-transformed epithelial cells examined have viability &#60; 10% under these conditions15,24. Thus, while there is a range in FSS resistance observed, the majority of the cancer cell lines tested exhibit greater FSS resistance than non-transformed cells. Cancer cell lines can be derived from both primary tumor tissues and metastases. One could postulate that cells derived from metastases may exhibit greater FSS resistance as this phenotype may have been selected during metastatic dissemination. However, the FSS resistance level was shown to not depend on whether cells were derived from primary tumors or metastases15,24. Moreover, the levels of FSS resistance did not correlate with metastatic potential in a series of human prostate cancer cell lines15.
To test this further, BALB/c mammary epithelial cells with varying metastatic potential (4T1 = highly metastatic, 4T07 = weak to moderate metastatic potential, 67NR = no to low metastatic potential25,26) were used. This experiment revealed that FSS resistance is not correlated with metastatic potential (Figure 1). Moreover, both 4T1 and 4T07 cells exhibit a biphasic loss of cell viability-a greater loss of viability in pulses 1-2 than observed in subsequent pulses. This is typical of most cancer cell lines investigated by this group. In contrast, 67NR exhibits a more linear loss of cell viability as a function of FSS. Collectively, the data from Table 2 and Figure 1 demonstrate that FSS resistance is a property of transformed cells.
<img alt="Figure 1" class="xfigimg" src="/files/ftp_upload/62478/62478fig01.jpg" /> 
Figure 1: Fluid shear stress resistance of syngeneic BALB/c mammary epithelial cancer cells. Cells were exposed to FSS (30 G needle, 10 pulses@250 mL/s), and viability was measured using resazurin conversion (n = 4/cell line). While FSS exposure reduced the number of viable cells (p &#60; 0.0001, 2-way ANOVA), and each cell line displayed different resistance profiles (p = 0.0446, 2-way ANOVA), there was no significant difference among cell lines after 10 pulses of FSS exposure (p = 0.2833, 2-way ANOVA). Abbreviations: FSS = fluid shear stress; ANOVA = analysis of variance. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/62478/622478fig01large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td colspan="2">Shear (&#964;):</td>
			<td colspan="3">wall (maximum)</td>
			<td colspan="9">minimum</td>
		</tr>
		<tr>
			<td colspan="2">Cell Diameter:</td>
			<td colspan="3">N/A</td>
			<td colspan="3">10 &#181;m</td>
			<td colspan="3">15 &#181;m</td>
			<td colspan="3">20 &#181;m</td>
		</tr>
		<tr>
			<td colspan="2">Needle Gauge:</td>
			<td>30</td>
			<td>27</td>
			<td>25</td>
			<td>30</td>
			<td>27</td>
			<td>25</td>
			<td>30</td>
			<td>27</td>
			<td>25</td>
			<td>30</td>
			<td>27</td>
			<td>25</td>
		</tr>
		<tr>
			<td rowspan="6">Flow Rate (&#181;L/s)</td>
			<td>20</td>
			<td>507</td>
			<td>220</td>
			<td>116</td>
			<td>32</td>
			<td>10</td>
			<td>4</td>
			<td>48</td>
			<td>16</td>
			<td>7</td>
			<td>64</td>
			<td>21</td>
			<td>9</td>
		</tr>
		<tr>
			<td>50</td>
			<td>1267</td>
			<td>550</td>
			<td>290</td>
			<td>80</td>
			<td>26</td>
			<td>11</td>
			<td>120</td>
			<td>39</td>
			<td>17</td>
			<td>159</td>
			<td>52</td>
			<td>22</td>
		</tr>
		<tr>
			<td>100</td>
			<td>2534</td>
			<td>1100</td>
			<td>580</td>
			<td>159</td>
			<td>52</td>
			<td>22</td>
			<td>239</td>
			<td>79</td>
			<td>33</td>
			<td>319</td>
			<td>105</td>
			<td>45</td>
		</tr>
		<tr>
			<td>150</td>
			<td>3801</td>
			<td>1650</td>
			<td>869</td>
			<td>239</td>
			<td>79</td>
			<td>33</td>
			<td>359</td>
			<td>118</td>
			<td>50</td>
			<td>478</td>
			<td>157</td>
			<td>67</td>
		</tr>
		<tr>
			<td>200</td>
			<td>5068</td>
			<td>2200</td>
			<td>1159</td>
			<td>319</td>
			<td>105</td>
			<td>45</td>
			<td>478</td>
			<td>157</td>
			<td>67</td>
			<td>637</td>
			<td>210</td>
			<td>89</td>
		</tr>
		<tr>
			<td>250</td>
			<td>6335</td>
			<td>2750</td>
			<td>1449</td>
			<td>398</td>
			<td>131</td>
			<td>56</td>
			<td>598</td>
			<td>196</td>
			<td>84</td>
			<td>797</td>
			<td>262</td>
			<td>111</td>
		</tr>
	</tbody>
</table>
Table 1: Maximum shear stress (&#964;wall) levels. The table lists the maximum wall FSS levels in dyn/cm2 for 30 G, 27 G, and 25 G needles at the flow rates of 20, 50, 100, 150, 200, and 250 &#181;L/s. Shear stress levels were calculated using the Poiseuille equation (<img alt="Equation 1" src="/files/ftp_upload/62478/62478eq01.jpg" />), available information for the inner diameter of each needle gauge, as well as the assumption that &#181; = 0.01 dyn&#183;s/cm2. Minimum FSS levels for each size were calculated using <img alt="Equation 2" src="/files/ftp_upload/62478/62478eq02.jpg" /> wherein r is the radius of cell, and R is the radius of the needle. Abbreviation: FSS = fluid shear stress; &#964; = shear; &#964;wall = maximum shear; &#181; = viscosity; Q = volumetric flow rate.
<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td>Cell Line</td>
			<td>Tissue Source</td>
			<td>Species</td>
			<td>Mean Viability (%) after 10 pulses</td>
		</tr>
		<tr>
			<td>TRAMPC1</td>
			<td>Prostate</td>
			<td>Mouse</td>
			<td>40</td>
		</tr>
		<tr>
			<td>4T01</td>
			<td>Breast</td>
			<td>Mouse</td>
			<td>32</td>
		</tr>
		<tr>
			<td>4T7</td>
			<td>Breast</td>
			<td>Mouse</td>
			<td>43</td>
		</tr>
		<tr>
			<td>67NR</td>
			<td>Breast</td>
			<td>Mouse</td>
			<td>46</td>
		</tr>
		<tr>
			<td>66CL4</td>
			<td>Breast</td>
			<td>Mouse</td>
			<td>28</td>
		</tr>
		<tr>
			<td>RT4</td>
			<td>Bladder</td>
			<td>Human</td>
			<td>62</td>
		</tr>
		<tr>
			<td>W17-266-4</td>
			<td>Melanoma</td>
			<td>Human</td>
			<td>46</td>
		</tr>
		<tr>
			<td>HS852</td>
			<td>Melanoma</td>
			<td>Human</td>
			<td>41</td>
		</tr>
		<tr>
			<td>HS695</td>
			<td>Melanoma</td>
			<td>Human</td>
			<td>41</td>
		</tr>
		<tr>
			<td>A2058</td>
			<td>Melanoma</td>
			<td>Human</td>
			<td>37</td>
		</tr>
		<tr>
			<td>A375</td>
			<td>Melanoma</td>
			<td>Human</td>
			<td>37</td>
		</tr>
		<tr>
			<td>RPMI-7951</td>
			<td>Melanoma</td>
			<td>Human</td>
			<td>35</td>
		</tr>
		<tr>
			<td>SKMEL2</td>
			<td>Melanoma</td>
			<td>Human</td>
			<td>29</td>
		</tr>
		<tr>
			<td>A101D</td>
			<td>Melanoma</td>
			<td>Human</td>
			<td>28</td>
		</tr>
		<tr>
			<td>MiaPaCa</td>
			<td>Pancreatic</td>
			<td>Human</td>
			<td>7</td>
		</tr>
	</tbody>
</table>
Table 2: Fluid shear stress resistance of various cancer cell lines. Each cancer cell line was exposed to fluid shear stress from the syringe and needle model (30 G needle, 10 pulses@250 mL/s) (n &#8805; 3/cell line), and viability was measured either by luciferase activity or resazurin conversion.

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

Nanyang Technological University

Research

JoVE Journal

Cancer Research

2.5K Views.  University of Iowa. Dit protocol stelt kankercellen in een suspensie bloot aan korte pulsen van vloeistofschaarspanning om bepaalde aspecten na te bootsen van hoe gemetastaseerde kankercellen worden blootgesteld aan hemodynamische krachten terwijl ze zich in de bloedsomloop bevinden. Het is een relatief eenvoudige techniek om korte pulsen van hoge vloeistofschaarspanning toe te passen. Let bij het proberen van dit protocol op ongeappte naalden en buig ze niet, omdat dit de toegepaste krachtschaarspanning zal ver...

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

Microfluidic Device for Recreating a Tumor Microenvironment in Vitro

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

Microfluidic Device for Recreating a Tumor Microenvironment in Vitro

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

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

Bioengineering

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 ...

Rapid Isolation of Viable Circulating Tumor Cells from Patient Blood Samples

Circulating tumor cells (CTC) are cells that disseminate from a primary tumor throughout the circulatory system and that can ultimately form secondary tumors at distant sites. CTC count can be used to follow disease progression based on the correlation between CTC concentration in blood and disease severity1. As a treatment tool, CTC could be studied in the laboratory to develop personalized therapies. To this end, CTC isolation must cause no cellular damage, and contamination by other cell types, particularly leukocytes, must be avoided as much as possible2. Many of the current techniques, including the sole FDA-approved device for CTC enumeration, destroy CTC as part of the isolation process (for more information see Ref. 2). A microfluidic device to capture viable CTC is described, consisting of a surface functionalized with E-selectin glycoprotein in addition to antibodies against epithelial markers3. To enhance device performance a nanoparticle coating was applied consisting of halloysite nanotubes, an aluminosilicate nanoparticle harvested from clay4. The E-selectin molecules provide a means to capture fast moving CTC that are pumped through the device, lending an advantage over alternative microfluidic devices wherein longer processing times are necessary to provide target cells with sufficient time to interact with a surface. The antibodies to epithelial targets provide CTC-specificity to the device, as well as provide a readily adjustable parameter to tune isolation. Finally, the halloysite nanotube coating allows significantly enhanced isolation compared to other techniques by helping to capture fast moving cells, providing increased surface area for protein adsorption, and repelling contaminating leukocytes3,4. This device is produced by a straightforward technique using off-the-shelf materials, and has been successfully used to capture cancer cells from the blood of metastatic cancer patients. Captured cells are maintained for up to 15 days in culture following isolation, and these samples typically consist of &gt;50% viable primary cancer cells from each patient. This device has been used to capture viable CTC from both diluted whole blood and buffy coat samples. Ultimately, we present a technique with functionality in a clinical setting to develop personalized cancer therapies.

Circulating tumor cells are isolated from the blood of cancer patients without inflicting cellular damage. Isolation of tumor cells is accomplished using a bimolecular surface of E-selectin in addition to antibodies against epithelial markers. A nanotube coating specifically promotes cancer cell adhesion resulting in high capture purities.

Circulating tumor cells (CTC) are cells that disseminate from a primary tumor throughout the circulatory system and that can ultimately form secondary ...

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 ...

Isolation of Proximal Fluids to Investigate the Tumor Microenvironment of Pancreatic Adenocarcinoma

Pancreatic adenocarcinoma (PDAC) is the fourth leading cause of cancer-related death, and soon to become the second. There is an urgent need of variables associated to specific pancreatic pathologies to help preoperative differential diagnosis and patient profiling. Pancreatic juice is a relatively unexplored body fluid, which, due to its close proximity to the tumor site, reflects changes in the surrounding tissue. Here we describe in detail the intraoperative collection procedure. Unfortunately, translating pancreatic juice collection to murine models of PDAC, to perform mechanistic studies, is technically very challenging. Tumor interstitial fluid (TIF) is the extracellular fluid, outside blood and plasma, which bathes tumor and stromal cells. Similarly to pancreatic juice, for its property to collect and concentrate molecules that are found diluted in plasma, TIF can be exploited as an indicator of microenvironmental alterations and as a valuable source of disease-associated biomarkers. Since TIF is not readily accessible, various techniques have been proposed for its isolation. We describe here two simple and technically undemanding methods for its isolation: tissue centrifugation and tissue elution.

Pancreatic juice is a precious source of biomarkers for human pancreatic cancer. We describe here a method for intraoperative collection procedure. To overcome the challenge of adopting this procedure in murine models, we suggest an alternative sample, tumor interstitial fluid, and describe here two protocols for its isolation.

Pancreatic adenocarcinoma (PDAC) is the fourth leading cause of cancer-related death, and soon to become the second. There is an urgent need of variables ...

Circulating Tumor Cell Lines: an Innovative Tool for Fundamental and Translational Research

Metastasis is a leading cause of cancer death. Despite improvements in treatment strategies, metastatic cancer has a poor prognosis. We thus face an urgent need to understand the mechanisms behind metastasis development, and thus to propose efficient treatments for advanced cancer. Metastatic cancers are hard to treat, as biopsies are invasive and inaccessible. Recently, there has been considerable interest in liquid biopsies including both cell-free circulating deoxyribonucleic acid (DNA) and circulating tumor cells from peripheral blood and we have established several circulating tumor cell lines from metastatic colorectal cancer patients to participate in their characterization. Indeed, to functionally characterize these rare and poorly described cells, the crucial step is to expand them. Once established, circulating tumor cell (CTC) lines can then be cultured in suspension or adherent conditions. At the molecular level, CTC lines can be further used to assess the expression of specific markers of interest (such as differentiation, epithelial or cancer stem cells) by immunofluorescence or cytometry analysis. In addition, CTC lines can be used to assess drug sensitivity to gold-standard chemotherapies as well as to targeted therapies. The ability of CTC lines to initiate tumors can also be tested by subcutaneous injection of CTCs in immunodeficient mice.
Finally, it is possible to test the role of specific genes of interest that might be involved in cancer dissemination by editing CTC genes, by short hairpin ribonucleic acid (shRNA) or Crispr/Cas9. Modified CTCs can thus be injected into immunodeficient mouse spleens, to experimentally mimic part of the metastatic development process in vivo.
In conclusion, CTC lines are a precious tool for future research and for personalized medicine, where they will allow prediction of treatment efficiency using the very cells that are originally responsible for metastasis.

Culturing CTCs allows a deeper functional characterization of cancer, through assaying specific marker expression, and assessing drug resistance and the ability to colonize the liver among other possibilities. Overall, CTC culture could be a promising clinical tool for personalized medicine to improve patient outcome.

Metastasis is a leading cause of cancer death. Despite improvements in treatment strategies, metastatic cancer has a poor prognosis. We thus face an ...

A Mouse Tumor Model of Surgical Stress to Explore the Mechanisms of Postoperative Immunosuppression and Evaluate Novel Perioperative Immunotherapies

Surgical resection is an essential treatment for most cancer patients, but surgery induces dysfunction in the immune system and this has been linked to the development of metastatic disease in animal models and in cancer patients. Preclinical work from our group and others has demonstrated a profound suppression of innate immune function, specifically NK cells in the postoperative period and this plays a major role in the enhanced development of metastases following surgery. Relatively few animal studies and clinical trials have focused on characterizing and reversing the detrimental effects of cancer surgery. Using a rigorous animal model of spontaneously metastasizing tumors and surgical stress, the enhancement of cancer surgery on the development of lung metastases was demonstrated. In this model, 4T1 breast cancer cells are implanted in the mouse mammary fat pad. At day 14 post tumor implantation, a complete resection of the primary mammary tumor is performed in all animals. A subset of animals receives additional surgical stress in the form of an abdominal nephrectomy. At day 28, lung tumor nodules are quantified. When immunotherapy was given immediately preoperatively, a profound activation of immune cells which prevented the development of metastases following surgery was detected. While the 4T1 breast tumor surgery model allows for the simulation of the effects of abdominal surgical stress on tumor metastases, its applicability to other tumor types needs to be tested. The current challenge is to identify safe and promising immunotherapies in preclinical mouse models and to translate them into viable perioperative therapies to be given to cancer surgery patients to prevent the recurrence of metastatic disease.

A mouse tumor model of surgical stress is used to explore how postoperative immune suppression promotes metastatic disease and to evaluate immunostimulatory perioperative therapies.

Surgical resection is an essential treatment for most cancer patients, but surgery induces dysfunction in the immune system and this has been linked to ...

Medicine

Modeling Brain Metastasis by Internal Carotid Artery Injection of Cancer Cells

Brain metastasis is a cause of severe morbidity and mortality in cancer patients. Critical aspects of metastatic diseases, such as the complex neural microenvironment and stromal cell interaction, cannot be entirely replicated with in vitro assays; thus, animal models are critical for investigating and understanding the effects of therapeutic intervention. However, most brain tumor xenografting methods do not produce brain metastases consistently in terms of the time frame and tumor burden. Brain metastasis models generated by intracardiac injection of cancer cells can result in unintended extracranial tumor burden and lead to non-brain metastatic morbidity and mortality. Although intracranial injection of cancer cells can limit extracranial tumor formation, it has several caveats, such as the injected cells frequently form a singular tumor mass at the injection site, high leptomeningeal involvement, and damage to brain vasculature during needle penetration. This protocol describes a mouse model of brain metastasis generated by internal carotid artery injection. This method produces intracranial tumors consistently without the involvement of other organs, enabling the evaluation of therapeutic agents for brain metastasis.

Brain metastasis is a cause of severe morbidity and mortality in cancer patients. Most brain metastasis mouse models are complicated by systemic metastases confounding analysis of mortality and therapeutic intervention outcomes. Presented here is a protocol for internal carotid injection of cancer cells that produces consistent intracranial tumors with minimal systemic tumors.

Brain metastasis is a cause of severe morbidity and mortality in cancer patients. Critical aspects of metastatic diseases, such as the complex neural ...

Intravital Microscopy of Tumor-associated Vasculature Using Advanced Dorsal Skinfold Window Chambers on Transgenic Fluorescent Mice

Tumor and tumor vessel development, as well as tumor response to therapeutics, are highly dynamic biological processes. Histology provides static information and is often not sufficient for a correct interpretation. Intravital evaluation, in which a process is followed in time, provides extra and often unexpected information. With the creation of transgenic animals expressing cell-specific markers and live cell tracers, improvements to imaging equipment, and the development of several imaging chambers, intravital microscopy has become an important tool to better understand biological processes. This paper describes an experimental design for the investigation of tumor vessel development and of therapeutic effects in a spatial and temporal manner. Using this setup, the stage of vessel development, tip cell and lumen formation, blood flow, extravasation, an established vascular bed, and vascular destruction can be visualized and followed. Furthermore, therapeutic effects, intratumoral fate, and the localization of chemotherapeutic compounds can also be followed.

This protocol describes the intravital imaging of transgenic mice expressing cell-specific fluorescent markers. Intravital imaging provides a non-invasive method for high-resolution observations in living animals using implantable windows through which the microscopic visualization of processes is possible. This method is especially useful to study longitudinal processes.

Tumor and tumor vessel development, as well as tumor response to therapeutics, are highly dynamic biological processes. Histology provides static ...

Simulating the Influence of a Bipolar Radiofrequency Hyperthermia System on Tumors

A Computational Modeling Approach to Investigate the Influence of Hyperthermia on the Tumor Microenvironment

The biophysical properties of the tumor microenvironment differ substantially from normal tissues. A constellation of features, including decreased vascularity, lack of lymphatic drainage, and elevated interstitial pressure, diminishes the penetration of therapeutics into tumors. Local&#160;hyperthermia within the tumor can alter microenvironmental properties, such as interstitial fluid pressure, potentially leading to improvements in drug penetration. In this context, multi-physics computational models can provide insight into the interplay between the biophysical parameters within the tumor microenvironment and can guide the design and interpretation of experiments that test the bioeffects of local hyperthermia.
This paper describes a step-by-step workflow for a computational model coupling partial differential equations describing electrical current distribution, bioheat transfer, and fluid dynamics. The main objective is to study the effects of hyperthermia delivered by a bipolar radiofrequency device on the interstitial fluid pressure within the tumor.&#160;The system of mathematical expressions linking electrical current distribution, bioheat transfer, and interstitial fluid pressure is presented, emphasizing the changes in the distribution of the interstitial fluid pressure that could be induced by the thermal intervention.

Author Spotlight: Computing the Effects of a Local Radiofrequency Hyperthermia Intervention on Tumor Biomechanics

The article describes a protocol to simulate the transient temperature profiles and the coupled spatiotemporal variation of the interstitial fluid pressure following the heating delivered by a dipolar radiofrequency hyperthermia system. The protocol can be used to assess the response of biophysical parameters characterizing the tumor microenvironment to interventional hyperthermia techniques.

The biophysical properties of the tumor microenvironment differ substantially from normal tissues. A constellation of features, including decreased ...

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

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