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

Podsumowanie

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.

Streszczenie

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 (<1 dyne/cm²) and the highest levels present briefly as cells pass through the heart and around heart valves (>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.

Wprowadzenie

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 body^2,3. While en route to these sites, circulating tumor cells (CTCs) exist within a dynamic fluid microenvironment unlike that of their original primary tumor^3,4,5. 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 colonies^6,7,8. 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 µm in diameter), most CTCs become entrapped in capillary beds (6-8 µm in diameter) for variable lengths of time (s to days) where they may die, extravasate, or be displaced to the next capillary bed^8,9,10,11. 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 (<1 dyn/cm²) and the highest at regions around heart valves and atherosclerotic plaques (>500 dyn/cm²)⁵. Mean wall shear stress in arteries is 10-70 dyn/cm² and 1-6 dyn/cm² in veins^16,17.

In the heart, cells may be exposed to turbulent flows around valve leaflets where very high-level, but very short-duration FSS may be experienced^18,19. 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 cells^21,22,23. Multiple "pulses" 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.

Protokół

1. Cell preparation

1. Release cells from tissue culture dish when 70-90% confluent by following the recommended guidelines for the cell line in use.
   1. 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).
   2. Aspirate the PBS before adding 1 mL of 0.25% trypsin using manufacturer's protocol.
   3. 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.
2. Place the cell suspension into a conical tube.
3. Determine the cell concentration and total cell number.
4. Pellet cells by centrifugation (300 × g for 3 min), aspirate the supernatant, and resuspend cells in serum-free tissue culture medium to 5 × 10⁵ cells/mL.
   ​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.

2. Fluid shear stress exposure

1. 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).
2. Draw the cell suspension into a 5 mL syringe, and attach a 30 G ½" 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 µm). 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.
3. Run the syringe pump, and collect the sheared sample in the cut tube at an approximate 45° angle to reduce foaming. Collect a sample depending on the type of viability assay or downstream assay needs.
   1. 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.
4. Draw the sheared suspension back into the syringe, carefully reattach the needle using pliers, and place it back into the syringe pump.
5. Repeat steps 2.3 and 2.4 until the cell suspension has been exposed to the desired number of pulses of FSS.
   ​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.

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.

1. For all measures of viability, collect a sample prior to exposing cells to FSS.
   1. For enzymatic assays, take duplicate 100 µL aliquots and place them into a 96-well plate.
   2. For flow cytometry, take one 500 µL aliquot and place it into a 1.5 mL tube.
   3. For clonogenic assay, collect a 100 µL aliquot.
2. Enzymatic assay
   1. Collect 100 µL samples after 1, 2, 4, 6, 8, and 10 pulses of FSS exposure and place them in a 96-well plate.
   2. Add the desired substrate, and follow the protocol for the assay used:
      1. For resazurin, add 20 µL of a 0.15 mg/mL solution to each well. Add 20 µL of 0.15 mg/mL resazurin solution to wells containing 100 µL of medium alone. Incubate for 2 h in a 37 °C tissue culture incubator. Measure the absorbance using a plate reader capable of reading fluorescence (579 excitation/ 584 emission).
      2. For luciferase-expressing cells, add 100 µL of 15 mg/mL D-luciferin to 5 mL of medium. Add 100 µL of that solution to each well containing cells. Wait for 5 min, and then read the plate using a reader compatible with luminescence.
      3. For WST-1, add 10 µL 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.
   3. Compare the averaged signal from each of the FSS-exposed samples to the averaged static control sample to obtain the percentage of viable cells.
3. Flow cytometry²⁴
   1. Collect 500 µL samples and place them into 1.5 mL centrifuge tubes after 1, 2, 5, and 10 pulses of FSS.
   2. Centrifuge samples (500 × g for 3 min), and discard the supernatants.
   3. Resuspend the pellets with 1 mL of calcium- and magnesium-free PBS, and centrifuge the samples (300 × g for 3 min).
   4. Suspend the pellets with 500 µL of fluorescence-activated cell sorting (FACS) buffer (PBS with 0.5% bovine serum albumin and 0.1% sodium azide) with counting beads and membrane-impermeable or viability dyes such as propidium iodide (1.75 µg/mL).
   5. Determine the viability by comparing the ratio of viable cells, normalized to counting beads, in sheared samples to that of the static sample.
4. Clonogenic assay
   1. Take 100 µL of the static sample, and add 900 µL of growth medium to make a 1:10 dilution.
   2. Take 100 µL of the 1:10 diluted sample, and add 900 µL of growth medium to make a final 1:100 dilution.
   3. Add 100 µL of the 1:100 dilution sample into each of 3 wells of a 6-well dish containing 2 mL of growth medium.
   4. Repeat steps 3.4.1-3.4.3 with samples that have been subjected to 10 pulses of FSS.
   5. Let the cells grow for 7-10 days without changing the medium, and check for colony formation. Once colonies of ≥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
   6. 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.
   7. Rinse with an excess of water, and let the plate dry
   8. Count the colonies (clusters of ≥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.

Wyniki

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 comparators^15,24. 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 ≥ 20% after 10 pulses of FSS at 25...

Dyskusje

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 cells^15,22,24. 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...

Materiały

Name	Company	Catalog Number	Comments
0.25% Trypsin	Gibco	25200-056
14 mL round bottom tubes	Falcon - Corning	352059
30 G 1/2" Needle	BD	305106
5 mL syringe	BD	309646
96-well black bottom plate	Costar - Corning	3915
Bioluminescence detector	AMI	AMI HTX
BSA, Fraction V	Sigma	10735086001
Cell Titer Blue	Promega	G8081
crystal violet	Sigma	C0775
D-luciferin	GoldBio	D-LUCK
DMEM	Gibco	11965-092
FBS	Atlanta Biologicals	S11150
PBS	Gibco	10010023
Plate Reader	BioTek	Synergy HT
Sodium Azide (NaN3)	Sigma	S2002
Syringe Pump	Harvard Apparatus	70-3005