the benefits of microfluidics for imaging cell migration

The benefits of microfluidics for imaging cell migration

L.L. Soon1, F. Braet1, K.R. Ratinac

1, M. Schuliga

2, H.-Y. Chien

1 and A. Stewart

2.

1Australian Centre for Microscopy and Microanalysis (ACMM), The University of Sydney, NSW 2006, Australia 2Department of Pharmacology, University of Melbourne, Victoria 3010, Australia

This review explores methodologies for in vitro imaging of microfluidics-driven cell migration. Cell migration is a

dynamic process in which cells move on top of other cells or inside matrices and enter blood vessels or lymph nodes to

reach specific destinations, such as the lungs or bone. Examples of cell migration in vivo include the homing of cancer

cells; the transformation and movement of lung cells under asthmatic conditions; the migration of somites during limb

formation; and the responses of the immune system to injury. Chemotaxis is the process in which cells react to

extracellular signals in a directional manner to reach target sites. Most in vitro studies, however, lack spatial discrimination

of signals and cannot induce cells to migrate directionally. While the homogeneous cell-culture conditions of in vitro

studies have provided most of our knowledge of cell signalling and behaviours, our understanding of such processes in

physiological conditions that involve varied microenvironments, such as the presence of growth-factor gradients, remains

relatively poor. Microscopy plays an important role in the recording of the dynamic changes that occur as cells migrate.

Imaging conditions to be considered include temperature, pH balance, multipositioning systems and optics. In this review,

we describe the advantages and inherent limitations of several live-cell imaging systems, as well as their consequences for

data interpretation and for our understanding of the cell biology of migration. We also explain how adding microfluidics to

microscopy can be used to produce localised signals that draw cells towards regions of highest signal densities. This

technological combination will be critical to unraveling the complex molecular mechanisms of cell chemotaxis.

Keywords Cell migration; live-cell imaging; microfluidics

1. Introduction

In multicellular organisms, the highly organised and stabilised cytoarchitecture of internal tissues is important for

normal organ functions and for survival. However, in certain developmental, physiological and disease conditions, the

integrity of cellular organisation is compromised producing migratory cells. A process that exemplifies this upheaval is

the epithelial–mesenchymal transition (EMT). During EMT, epithelial cells, which are cuboidal in shape and line the

walls of luminal cavities, lose their adhesion to other cells and to the basement membrane and thereby transform into

small groups of cells or individual stellate-shaped (i.e., star-like) mesenchymal cells. Furthermore, in diseases such as

cancer, the affected epithelium no longer functions properly in monitoring and facilitating the secretion and absorption

of macromolecules, while the untethered cells become proliferative and/or motile. These effects can be devastating if

the cells multiply to form a primary tumour, disperse to colonise distant sites and form secondary tumours. In asthmatic

patients, the accumulation of mesenchymal cells, such as fibroblasts and myofibroblasts, in the airway wall is possibly

due to the migration of cells via the circulatory system 1,2 and from pre-existing sub-epithelial fibroblasts, and/or EMT

3.

The consequences of increased mesenchymal-cell burden are elevated matrix deposition and its compaction, leading to

fibrosis or stiffening and thickening of the airway walls, that are amongst the hallmarks of asthma. During limb

formation in the developing embryo, skeletal-muscle-progenitor cells migrate to the limbs, the tongue and the

diaphragm, where they differentiate to form skeletal muscle. These cells undergo EMT prior to delaminating from the

dermomyotome, an epithelial structure, and migrating to populate destination sites 4. Apart from cells originating from

the epithelia and stroma, there are circulating and tissue-resident immune cells capable of efficient, long-distance

migration to monitor and protect the body from pathogens and foreign material.

Clearly, migration of cells to target sites or chemotaxis towards regions of high ligand density is integral to many

physiological and pathophysiological (i.e., disease) processes. Understanding the fundamental principles and

determining the molecular mechanisms involved are critical for continuing to advance cell biology and for developing

counter measures to eradicate migration-related diseases.

2. Cell migration

While EMT is an important process in embryonic development and in many diseases, its role in cancer is among the

most widely studied of such phenomena. Given the importance of this field to biology and medicine, as well as our

interests in the area, we will focus here only the migration behaviour of cancer cells.

During EMT in cancer, epithelial cells undergo changes that result in the dissolution of an intact epithelium through

disruption of cytoskeletal organisations 5, of membrane specialisations

6, and of cell-to-matrix and cell-to-cell adhesion.

This creates at least three types of migratory tumour cells: cells that move as a cohort; single cells that display

mesenchymal characteristics; and single cells that exhibit amoeboid characteristics. Cohort cells migrate as a coherent

Microscopy: Science, Technology, Applications and Education A. Méndez-Vilas and J. Díaz (Eds.)

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group in which the central cells are organised in a sheet and are cuboidal in shape, while the leading cells are more

mesenchymal-like in appearance, with an elongated stellate shape 7. Singly migrating, mesenchymal-like tumour cells

are highly polarised with leading protrusions and a long trailing-edge. These cells have the appearance of, and migrate

like, normal mesenchymal cells such as fibroblasts 8. In contrast, amoeboid-like tumour cells are not angular, have

lower intrinsic cell polarity, and demonstrate loss of stress fibres and focal contacts 9,10

. These cells display morphology

and movement similar to that of classical amoeboid cells such as neutrophils, which are a type of white blood cells, and

the cells of the amoeba, Dictyostelium. Despite having acquired characteristics akin to normal chemotactic cells, all

types of tumour cells typically move 10–15 times more slowly than neutrophils or Dictyostelium cells. Another distinct

difference between amoeboid-like tumour cells and normal amoeboid cells is the tumour cells’ inability to maintain

persistent movement under a uniform concentration of chemoattractants (i.e., in the absence of chemical gradients). In

such an environment, the tumour cells make numerous changes in the direction of movement and so demonstrate

random walking 11. Neutrophils and Dictyostelium cells, on the other hand, adopt a protrusion in a stochastic manner in

an arbitrary direction, and then maintain that asymmetry and show persistent movement in that direction, which results

in so-called linear walking 12. Genuine amoeboid cells, therefore, exhibit internal polarity that is lacking in their

amoeboid tumour-cell counterparts. One advantage of such internal polarity could be that, once a direction of

movement is determined, marshalling of receptors to the leading-edge increases sensitivity to shallow chemoattractant

gradients or weak chemoattractant levels; another could be that orientation cues are recorded and retained for longer

periods.

Other cellular alterations that occur during EMT include aberrant autoactivation of receptors. In the normal and intact

epithelium, the apical surface is directed towards the lumen where ligands to receptors are secreted. However, the

corresponding receptors are found in the basolateral region of epithelial cells, which effectively isolates the receptors

from the ligands. EMT results in loss of epithelial polarity and the onset of autocrine signaling whereby the tumour cells

can self-activate surface receptors with secreted ligands 13. In addition, surface receptors are redistributed globally on

the cell membrane in cancer cells, with the consequence of reducing localised sensitivity to ligands. Often, the reduction

in receptor density is compensated by the increased expression of growth-factor receptors that commonly occurs during

carcinogenesis. High levels of receptor expression, however, can lead to receptor autoactivation 14,

15 and/or signal

propagation 16 due to the existence of a threshold receptor density, creating a bistable activation regime. Thus, steady-

state receptor-activation conditions that occur independently of ligands can exacerbate the loss of sensitivity to external

directional cues in tumour cells.

Amoeboid tumour cells, therefore, must strike a fine balance between achieving greater surface area that is receptive

to chemoattractants and maintaining sensitivity to external growth-factor gradients. As explained above, this sensitivity

is greatly reduced due to global redistribution of receptors, due to autocrine signaling and due to receptor autoactivation,

resulting in random motility 17. Amoeboid-like tumour cells appear to have developed adaptive responses to enable

efficient chemotaxis; these include paracrine interactions with macrophages at short-range through cross-receptor

activation by secreted ligands 18, and proteolysis of ECM fragments to produce EGF-like ligands that cause a local

gradients of chemoattractant 19.

It should be clear by now that tumour cells differ from each other and from normal cells in their intrinsic polarities,

which means they require different external gradients to chemotax. Consequently, one of our endeavours has been to

create steep chemoattractant gradients that we hypothesise will be more suitable for inducing tumour-cell chemotaxis 20,21,22

. Characterisation of the migration properties of tumour cells will shed light on the diversity of regulatory

mechanisms involved and, at the same time, highlight shared principles that underlie the biology of chemotaxis. It is, of

course, equally important in working towards viable medical treatments that will halt, or even prevent, the spread of

cancer in patients.

3. Microscopy

When it comes to imaging the migration of cells, three main factors will determine the success of the experiment: the

type microscopy selected, the image-capture technology used, and the type of chemotaxis chamber employed. Each is

important in its own right; yet one cannot be selected in isolation from the others. In this section, we will critically

consider microscope and detector options. Chamber design is addressed in the next section.

3.1. Live-cell imaging

Many different microscopy techniques can be used for viewing cell migration, including transmitted-light, wide-field,

confocal, multiphoton, and ultra-resolution techniques. However, the most appropriate technique depends on the chosen

application and on the construction of the chemotaxis chamber. Here, we focus on the strengths and weaknesses of

some of the microscopes used in imaging cell migration, with some emphasis on the detection techniques. These have

undergone some revolutionary changes in the recent past, but have not yet been widely implemented.

Imaging imposes stress on biological systems that can cause the deterioration of cells very rapidly. To maintain cell

viability, the environment of the cells is kept under near-optimal conditions during imaging. A microscope enclosure


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equipped with heating reduces thermal fluctuations. Objective lenses and the microscope stage can also be heated

separately, thereby minimising the lost of heat. Carbon dioxide (CO2, 5%) and humidity (>88%) controls can be

installed to mimic the slightly acidic and humidified in vivo conditions. Alternatively, buffered isotonic media, such as

HEPES made to 20 mM and pH 7.5 or the commercially available medium Lebovitz (L15, Invitrogen) can be used in

the absence of CO2 incubators. These media will maintain cell physiology for a couple of hours during imaging. For

general measurements of movement parameters, digital image contrast (DIC) or phase-contrast microscopy with a 40x

objective is adequate. These techniques use optical insertions in the microscope light-path to create differentials in the

resulting waveforms, providing greater contrast for thin, transparent specimens such as cells. Time-lapsed series of

varying intervals will generate movies that can be analysed offline.

For the study of protein dynamics or structural changes during cell migration, fluorescence or confocal microscopy

of cells tagged with live probes offers greater specificity than transmitted-light microscopy. Fluorescent protein (FP)-

tagged proteins of interest are preferably expressed by native promoters to allow physiological levels of expression,

thereby avoiding overly high expression levels that can lead to non-specific localisation, to congestion in trafficking

routes and/or to decreased cell viability. GFP-actin expression driven by the actin promoter, for example, has been used

to understand actin dynamics during cell migration as well as serving as a general cell marker that defines the cell form,

which is ideal for tracking studies. The GFP-actin molecules also clearly demarcate filopodia and lamellipodia,

allowing dynamic studies of the formation of these migration structures.

For other studies, spatial resolution is more critical than speed and, for these applications, wide-field microscopy is

often unsuitable, as the images suffer from far greater degradation of the lateral and axial point spread functions than

occurs in confocal microscopy. Multiphoton microscopy has the added advantage over other fluorescence techniques of

allowing deep sectioning into samples, reaching up to one millimetre in depth, surpassing that of confocal microscopy.

Consequently, multiphoton microscopy is suitable for viewing thicker samples, tissues and organs 23. It offers similar,

or possibly slightly worse, resolution than confocal microscopy due to the use of near-infrared wavelengths of light as

the excitation source. The spinning-disc confocal microscope, which collects light from parallel arrays of pinholes,

exceeds the speed of all point-scanning instruments. The instrument can, in theory, scan up to 1000 frames per second

(fps) compared with 10-20 fps for a confocal microscope; however, the spinning-disc instrument approaches confocal

resolutions for thin specimens only. The Yokogawa system, which incorporates microlenses to focus light onto the

pinholes, is an innovation that significantly improves light collection, gathering up to 60% of all fluorescence emitted

from samples. The Yokogawa spinning-disc confocal microscope coupled with an electron-multiplying charge coupled

device (EM-CCD) camera (see Section 3.2 below), is the ideal system for live-cell imaging because of excellent light-

gathering ability, low phototoxicity and high speeds in image capture and light detection.

In terms of phototoxicity, there are advantages and disadvantages associated with the various fluorescence techniques

and the excitation wavelengths used. While longer wavelengths in the red and near-infrared region of the

electromagnetic spectrum might be gentler on the cells, the extra heat generated can damage cells. Therefore, IR filters

should be used to minimise heat transfer to the cells. At the other extreme, UV light is deleterious to cells, as it tends to

damage DNA, and should be avoided if possible. Cells in general, react adversely and tend to move away from all

wavelengths of light, including white light. Therefore, neutral density filters should be in place to reduce light intensity.

Other precautions include the installation of shutters, minimising exposure to fluorescence light by first focusing with

transmitted light, avoiding loss of light by removing DIC or phase optics, and reducing exposure times to the minimum

for obtaining reasonable signal-to-noise ratios.

3.2. The detectors

The optimal detector can change according to the demands of the research, the microscope used and the properties of

the detector. Key among these are the frame rate and the quantum efficiency, which measures how well photons are

captured by the detector. Here we will provide an overview of the two main classes of light detectors, photomultiplier

tubes and charge coupled devices, and some advice on detector selection.

Photomultiplier tubes (PMTs) are the most commonly applied detectors in confocal microscopy. Photons enter the

PMT via a quartz window before reaching a photocathode. This causes the release of electrons that are, in turn, directed

towards a series of dynodes. The dynodes serve to multiply the number of electrons and the amplification can be varied

by tuning the voltage so that, for a 12–14 dynode PMT, high gains can be attained. An important consideration when

selecting a camera is the frame rate, which indicates the number of consecutive images produced per second and is

expressed either in units of fps or of Hertz (Hz). Unlike the case for charged coupled devices (CCDs; see below), the

electrons in PMTs do not dwell in pixel wells and the response to input photons occurs within nanoseconds, all of which

makes PMTs potentially the fastest light detectors. However, in confocal microscopy each frame is generated from a

series of point and line scans, so that the frequency of the scanner is a limiting factor in detection speed, in general. For

a 500 Hz scanner that acquires 500 lines per second, the frame rate is approximately 0.5 fps over a 1024 x 1024 image.

Of course, the time required for detection is far greater for live-cell imaging in three-dimensions, as each time frame

consists of a stack of two-dimensional images, and the stacks are connected through time for defined imaging periods.

Significant increases in detection speed can be achieved, but only at the cost of image resolution or one lateral

dimension, by capturing images of only 128 x 128 pixels or, where appropriate, by performing single-line scans.



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However, applications for these scanning parameters are limited, especially as image averaging is not an option during

high-speed imaging. Moreover, the quantum efficiency of most PMTs is less than 25%, meaning that 75% of photons

that reach the PMT are not detected. Therefore, where light levels are low or high sensitivity is required, PMTs are

probably not the best detector to use 24.

Charged coupled devices (CCDs) comprise a silicon wafer with an etched integrated circuit of light-sensitive

elements or pixels. Photons falling onto the doped-silicon surface generate charges that are then converted into voltage

or electronic signals. At its most basic, a CCD camera consists of photodiodes, a readout register, a horizontal serial

register and an on-chip floating diffusion amplifier. The photodiode accepts incoming photons that are simultaneously

shifted to the adjacent readout register. The horizontal rows of information are then transferred, line by line, to the

horizontal serial register, where they are converted into a voltage by the analog-to-digital electronics.

The quantum efficiency in CCDs measures the conversion of photons into photoelectrons (electric signal) in the CCD

pixels. In this case, the quantum efficiency is compromised by readout noise and dark current. The readout occurs at the

analog-to-digital transformation stage when accumulated charges are converted into current. Readout noise is generated

primarily when the amplifier is reset for the incoming pixel, after the accumulated charge in each pixel has been

converted into a voltage. It is also proportionately dependent on the pixel clocking frequency: the faster the pixel clock,

the greater the readout noise. However, increasing the readout speed allows higher gains to be reached, maximising

coverage across the dynamic range of the camera. Reducing the clocking speed will reduce readout noise, but could

compromise studies that require high-speed dynamic imaging. The other quantum-efficiency-limiting factor, dark

current, results from thermal migration and excitation of extraneous electrons into the image pixels. It is also known as

thermal charge and cannot be distinguished from the photoelectrons that contribute to the image. Dark current can be

reduced by decreasing the temperature of the device and, therefore, CCDs are typically cooled to between –30°C and –

100°C, which requires thermoisolation under vacuum.

The frame rate depends on the amount of input information or the number of pixels to be read, and the clocking

speed of the pixels in the floating diffusion amplifier. Decreasing the number of pixels or increasing the speed of the

pixel clock will increase the frame rate, but this could also result in more readout noise. Binning is a method of

combining signals (i.e., adding the stored charges) in a CCD that can be used to reduced the number of pixels to be

processed. Binning options such as 2x2, 4x4 and 8x8 are commonly used to improve signal-to-noise ratio, and to

increase sensitivity and the frame rate; however, all of these benefits come at the expense of image resolution.

Improvements to CCDs include the development of back-illumination, in which excess silicon is etched away on the

bottom of the chip to allow photons to pass through the back of the device, rather than through the gate structure on the

top of the CCD. This significantly increases photon access to the pixel wells, improving the quantum efficiency and

dynamic range of the sensor. Front-illuminated CCDs, for example, have poor detection in the blue wavelength range of

400–500 nm, whereas these photons can be easily detected by back-illuminated CCDs.

The EM-CCD is another innovation that has increased sensitivity and readout speed without an increase in the

readout noise. The technology involves the integration of an electron multiplier into the device. The multiplier increases

the electron counts in the serial register, prior to conversion into a voltage by the floating diffusion amplifier. This

multiplication of weak signals at the early stage prevents additional increases in readout noise due to the resetting of the

amplifier. Therefore, high quantum efficiencies can be achieved, even at high clocking speeds, with negligible readout

noise. The gain register is adjustable from the software and weak signals can be tweaked for visibility above the readout

noise at any clocking speed. The EM-CCD technology has therefore overcome an intrinsic weakness in conventional

CCD, in which speed and sensitivity are incompatible, and has enabled greater sensitivity without compromising speed.

An EM-CCD system combines cooling to suppress dark current, a gain register that minimises readout noise and back-

illumination, all of which results in high quantum efficiency and sensitivity. Indeed, the EM-CCD camera is capable of

imaging single-photon events without an image intensifier, allowing use of up to 95% of the the sensor’s dynamic

range.

Which detector to choose? EM-CCDs have been designed for capturing weak signals and for dynamic applications.

Therefore, this is the ideal choice for live-cell imaging where low light levels are required to preserve cell viability 25.

There is also the added flexibility of the tunable gain register for multiplication of signals in real-time, as needed. On

the other hand, if the signal is strong and well-above the readout noise and speed is not a critical factor, a high-quantum-

efficiency CCD can produce the greatest signal-to-noise ratio. The quantum efficiency of CCDs reaches between 60-

70%, achieving high resolutions of 0.1 µm pixels. This further allows binning or summing of pixels to increase signal-

to-noise with some sacrifice of the resolution. Therefore, applying longer exposures, slower readouts and binning in a

CCD can produce high-quality images with excellent signal-to-noise. This type of camera will be appropriate for

observations of morphological changes or actin-relation polymerisation of cell structures that occur in the timeframe of

hundreds of milliseconds to minutes 26.

4. Microfluidic devices

Originally, chemotaxis was observed with the Dunn 27 and Zigmond

28 chambers. The basic setup used a spezial

microscope slide with a raised platform to hold the cells, and an adjacent well of chemoattractant. When a coverslip was



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placed over the slide, the chemoattractant was drawn onto the viewing surface, producing a gradient that remained

stable enough for the analysis of chemotaxis 29,30,31

. In the absence of a flow system, the gradient would decay over one

to several hours.

The integration of microfluidics with dedicated viewing chambers can overcome limitations in gradient stability. The

micropipette assay, for example, was the prelude to microfabricated chambers with fluidic inlets and outlets. Designed

originally for the microinjection of particles into cells, the device consists of a motorised manipulator, which moves the

micropipette into position, a glass micropipette filled with solution and the injector that ejects the fluid. The

micromanipulator is used to position the micropipette approximately five micrometres away from the tumour cell. An

input pressure of approximately 30 Pa is sufficient to extrude the chemoattractant locally in the vicinity of the cell. The

tumour cells respond accordingly, projecting a cell protrusion in the direction of the gradient. However, due to the close

range of the micropipette, cell receptors quickly become saturated, preventing the cell from performing a full

chemotaxis cycle. Therefore, this assay is adequate only for studying the early stages of the process, such as sensing and

protrusion formation.

A modification of this assay serves to divert the flow of chemoattractant away from the cells and at the same time

develop a steep gradient sufficient to induce the chemotaxis of amoeboid tumour cells. This was achieved by

positioning the tip of the micropipette below the cell growing surface to produce a sharp and stable gradient at the cell

growing surface that induces the chemotaxis of cells (Fig. 1A).

Figure 1. Stylised chambers used in the live-viewing of cell chemotaxis. (A) This micropipette chamber consists of two coverslips

where the upper one is shorter than the lower one. A micropipette abuts the edge of the short coverslip, diverting the flow of

chemoattractant so that it rises vertically and spills over the upper coverslip, ‘wetting’ the edge to form a steep gradient. (B) This

vertical-aperture chamber consists of two transparent wafers. The surface of the lower wafer is etched with grooves that transforms

into microchannels when capped by the upper wafer. The upper wafer contains vertical grooves that connect the ends of the

microchannels to the surface of the cell chamber. Chemoattractant is conducted through the microchannels and released from the

apertures to form a gradient.

The use of microfluidics in the study of cell migration has entailed developing special devices that allow the flow of

fluids across the cell-growing surface. These devices consist of a transparent chamber, for growing and viewing cells, in

which inlets and outlets allow flow of external factors under pressure. In the absence of more sophisticated pump

systems, compressed air, from outlets commonly present in microscopy laboratories, is sufficient to generate the flow in

the microfluidic chambers (Fig. 1B). These chambers are made possible by adaptations of micro-electromechanical

systems (MEMs) technology originally developed for the miniaturisation of parts in electronic devices. In recent times,

the development of soft lithography techniques have led to greater flexibility in the conformations of the microelements 32.The attractions of this technology to biologists include the varied possibilities in the construction of celluar

environments 33,34,35

, the increase in throughput and conservation of time and reagents. The microfluidic devices

constructed for the study of chemotaxis consist of a viewing surface where cells are grown and apertures, either laterally 9,36,37

or vertically 20,22

placed, that extrude chemoattractants in nanolitre volumes. Laminar flow of the released

chemoattractants results in mixing with the surrounding media primarily by diffusion. The lack of turbulent flow

ensures gradients remain stable for long periods, typically up to several hours.

In a device where lateral apertures are used, there are two sources of fluids that flow into a series of parallel and anti-

parallel channels to allow mixing of the chemoattractants before they are finally released into the cell-growing chamber 38. In these assays, the cells are first released into the chambers and allowed to settle or adhere prior to the formation of

the chemoattractant gradient. Cell chemotaxis can then be imaged and analysed. A drawback to these devices is that the

flow of chemoattractant is directed towards the cells. At typical chemoattractant flow rates of millimetres per second,

the flow could exert shear stresses on cells, making this assay unsuitable for some cell types, such as mesenchymal-like

tumour cells that are susceptible to rupture in blood vessels 39. Further modifications of the chambers, such as the

introduction of matrices, have significantly slowed down the flow rates to more physiologically relevant levels of 0.6–

6.0 µL/min, avoiding shear stresses 38.

In a device where vertical apertures are present, chemoattractants are delivered by underlying microchannels and

released into the overlying cell-growing surface through vertical apertures (Figs. 1B, 2) 20. The tumour cells are grown



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in the cell chamber overnight, the stage is heated for at least two hours prior to the experiment and the media replaced

with supplemented L15 media prior to imaging. A time-lapsed sequence and the flow of chemoattractant are activated

to record chemotaxis. A stable and local gradient is produced that attract the migration of slow-moving cells such as

cancer cells (Figs. 2A,C) 40. Furthermore, by directing the flow of chemoattractant vertically, shear forces do not act on

the cells. The downsides to this method are that vertical apertures are difficult and expensive to produce, and that

vortices can form in the upper reaches of the gradient 20. Cells are, however, typically only five micrometres in height

with one-micrometre-thick lamellae and, therefore, respond well to the stable gradients present at the growth surface

(Fig. 2D). However, only cells lying closest to the gradient sources respond to the chemoattractant and cells further

behind do not chemotax (Fig. 2D). This result is accounted for by the preference of these cells for localised, steep

gradients; in an assay where the gradient is shallow but more widely distributed, these tumour cells will not chemotax

irrespective of where they are located relative to the gradient source.

Figure 2. A chemotaxis assay with vertical apertures. A. The chemotaxis chamber containing cells is placed on a heated stage of an

inverted microscope. The time-lapsed sequence is initiated prior to switching on the microfluidic flow. B. A segmented view of the

chamber which consists of a metal casing that can be tightened, a coverglass coated with a UV15 layer which contains the

microchannels, a capping layer which seals the microchannels and where 1 micron apertures are created at the ends of the

microchannels, a soft PDMS layer with large openings that form the walls of the cell chamber and chemoattranctant well and a hard

PMMA layer which helps to seal the chamber. C. Measurements of FITC-dextran used in a microfluidic flow experiment indicate

that a stable gradient exists for the duration of the chemotaxis assay. D. Towards the end of the assay, tumour cells (highlighted) have

moved to the source of chemoattractant.

More recently, MEMs and microfluidics technologies have been applied to 3D cellular environments to recreate the

more naturally diffusive, multidimensional in vivo tissue conditions 41,42,43,44

. Using soft lithography to generate silicon

molds containing raised relief structures, Cheng et al., 41 constructed a chamber consisting of 3 parallel channels within

a block of hydrogel. The first channel contains the chemoattractant whilst the middle channel has cells and the third

constitutes the output well. The gradient of chemoattractant that forms between the first and third channels allows the

chemotaxis of cells from the middle channel. The highly porous matrices can only sustain a shallow gradient, which is

sufficient for evaluating the chemotaxis of neutrophils and other cells. For example, the gradient across a single cell

located in the middle channel approximates 33% 41 compared with 68%-100% (Fig. 2C) required for the chemotaxis of

amoeboid tumour cells on 2D matrices. Three-dimensional chambers will nevertheless, become more important in the

future to simulate certain natural environments. Adaptation of the chambers to variable gradient needs of cells will be

necessary.

There are several important considerations when integrating microfluidics with live-cell imaging. Firstly, the active

surface of the device must be non-toxic to cells and the chamber must withstand wet conditions and temperatures of

37°C. Secondly, the configuration of the fluidic device needs to be able to be physically accommodated within limited

microscope-stage areas. Thirdly, the active surface of the device has to be compatible with microscope optics. The latter

is critical for resolving fine structures or detecting molecular events such as protein–protein interactions.

In studies not requiring live imaging, high numerical aperture (NA), oil-immersion objective lenses are suitable due

to the approximate compatibility of the lens, oil, embedding medium and coverslip. In live-imaging where air and



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aqueous fluid lie between the lens and the specimen, the accumulation of aberrations becomes problematic. The use of a

water-based objective lens significantly overcomes incompatibility in the elements lying in the light path. Another

consideration is the thickness of the coverslips, which have been standardised to allow for correction of image

aberrations during the design of lenses. Standard recommended coverslips (#1.5) have a thickness of approximately 1.7

mm. Thinner coverslips are also available (#0) with a thickness of 0.08–0.13 mm, and these can be used with objective

lenses that have a correction collar that moves internal lens elements to compensate for variations in coverslip

thicknesses. The #0 coverslips are suitable for imaging thicker samples such as tissue slices or gelled matrices.

If high resolution is needed, the microfluidic device must be constructed with a base or an active cell surface that is

transparent and that is optically compatible with the working distance of the objective lens. If the surface is too thick,

which is true for many fluidic or lab-on-a chip devices, then the working distance will be incorrect and the samples will

not be in focus. This problem is evident for objective lenses with higher numerical apertures, but objective lens with a

numerical aperture of 0.4 or lower is significantly more tolerant of thick active surfaces.

5. Conclusions

Long-term and short-term chemotaxis assays have led to greater understanding of the processes of cell migration.

Many of the migratory steps and mechanisms are universal among cell types. Where there have been differences, it has

turned out that the plasticity of cells dictates alternations between different mechanisms, suggesting a continuum of,

rather than distinct variances in, migratory styles. How cells sense the microenvironment is one example where a

universal concept applies, albeit that the molecular pathways and players vary across different cell types. The sensing

process is defined by the formation of an internal gradient of molecules that exceeds the strength of the external

gradient of factors 21,45,46,47

. The concept that describes how this happens is called ‘local activation and global

inactivation (via a diffusible inhibitor)’, a process in which a positive signal, such as active cofilin, is amplified at the

cell front while a negative regulator, such as LIM kinase, concentrates to the rear of amoeboid tumour cells. These

studies were mainly performed with the micropipette assay 48,49

and other local activation experiments using caged

molecules (of cofilin) 50,51

. Preceding studies, which were seminal to understanding sensing, demonstrated the

polarisation of PIP3 at the cell front and PTEN at the rear of Dictyostelium cells 52,53

and similar behaviour in

neutrophils 54.

In amoeboid tumour cells, it is clear that an internal gradient of cofilin activity is established during sensing as the

molecule severs actin polymers at the cell front, expanding the actin dendritic array and forming the leading protrusion.

However, it is unclear how the cells subsequently recover from the initiation of directional migration to continously

develop sensitivity to the external gradient. In addition, it is unknown whether the same molecules regulate sensing in

mesenchymal-like tumour cells, which have polar and cytoskeletal properties that are distinct from the amoeboid-like

cells.

There are many more questions to be asked and answered about the mechanisms of chemotaxis, which might be

widely applicable to different cell types. For example: how do adhesion receptors and growth-factor receptors direct

their many similar downstream signals into functional, dynamic networks in the polarised, motile cells? and how do

these different receptor types spatially and temporally segregate the activation of the same downstream effectors? High-

resolution microscopy, coupled to long-term migration assays in microfluidics-driven gradients, will provide the means

to address these queries and others. Even though these in vitro assays explore only one aspect of very complicated

internal environments, there is true value in being able to describe and generalise major processes of cell chemotaxis

that currently evade us. Unlocking chemotaxis mechanisms across different cell types, with functions in many

physiological processes as well as many diseases, will facilitate further advances in the life sciences, from the levels of

molecules to tissues.

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the benefits of microfluidics for imaging cell migration

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