PAPER
www.rsc.org/loc | Lab on a Chip
A simple microfluidic method to select, isolate, and manipulate single-cells in mechanical and biochemical assays† Sylvain Gabriele,a Marie Versaevel,a Pascal Preirab and Olivier Theodoly*b Received 2nd February 2010, Accepted 9th February 2010 First published as an Advance Article on the web 24th March 2010 DOI: 10.1039/c002257h This article describes a simple and low-tech microfluidic method for single-cell experimentation, which permits cell selection without stress, cell manipulation with fine control, and passive selfexclusion of all undesired super-micronic particles. The method requires only conventional soft lithography microfabrication techniques and is applicable to any microfluidic single-cell circuitry. The principle relies on a bypass plugged in parallel with a single-cell assay device and collecting 97% of the flow rate. Cell selection into the single cell device is performed by moving the cell of interest back and forth in the vicinity of the junction between the bypass and the analysis circuitry. Cell navigation is finely controlled by hydrostatic pressure via centimetre-scale actuation of external macroscopic reservoirs connected to the device. We provide successful examples of biomechanical and biochemical assays on living human leukocytes passing through 4 mm wide capillaries. The blebbing process dynamics are monitored by conventional 24 fps videomicroscopy and subcellular cytoskeleton organization is imaged by on-chip immunostaining.
1. Introduction There is an increasing interest from a variety of disciplines for single-cell experimentation methods,1 because they permit the study of cellular processes dynamically at the sub-cellular level. Significant cellular phenotypes have, for instance, been resolved by single-cell experiments,2–4 which could not have been detected on the average behaviour of a population. In this context, microfluidics is a method of choice for single-cell experimentation, because it permits integration of the selection, navigation and positioning of a single-cell5–13 with a large variety of analysis methods. Microfluidic cellular operations at the micrometre scale minimize sample contamination and product loss, which would preclude sensitive, reproducible, and quantitative single-cell analysis.14 Many efforts for single-cell manipulation have been driven by the goal of analyzing the content of individual cells (DNA and protein)15–18 in order to develop micro total analysis systems (m-TAS). These systems present specific challenges in terms of the detection of low-level signals with small amounts of analytes, whereas the main difficulty of single-cell analysis arises from the manipulation and sorting of micron-size objects with high precision in confined micrometre-sized environments. Several sophisticated technical solutions have been developed to a Universit e de Mons, Laboratoire Interfaces & Fluides Complexes, Centre d’Innovation et de Recherche en Mat eriaux (CIRMAP), 20, Place du Parc, B-7000 Mons, Belgique b Universit e de la M editerran ee, Adhesion & Inflammation, INSERM U600-CNRS UMR6212, 163 Av. de Luminy, F-13009 Marseille, France. E-mail:
[email protected]; Fax: +33 (0)4 91 82 88 69; Tel: +33 (0)4 91 82 88 51 † Electronic supplementary information (ESI) available: Movies showing: zero flow and flow conditions, cell selection, passage of a neutrophil in a 10 3 mm constriction, leukocyte THP-1 through a succession of 5 mm wide constrictions, doublet of cells in sequential constrictions. See DOI: 10.1039/c002257h
This journal is ª The Royal Society of Chemistry 2010
achieve these tasks. Multilayer soft lithography has been used to develop an integrated cell sorter with various functionalities (microvalves, dumps, micropumps, .) to perform cell sorting in a coordinated and automated fashion, by purely fluidic manipulation of cells.19 Electrokinetics,20–23 dielectrophoresis (DEP),9,24–29 and optical trapping5,30–32 have also been successfully used for manipulation and sorting of a wide variety of cells in microchannels. However, these techniques have some limitations or inconveniences. Electrokinetics, which is the most common technique for the control of flow in microfluidic devices,11 imposes high current and Joule heating, leading to potential cell damage or lysis.33 Most methods for cell control require multiple integrated pumps, sensitive optics and fast electronics, and thus, remain complex and expensive. Therefore, although these sophisticated technological achievements have made it possible to build automated analysis microfluidic systems for single-cell experiments, much lab-scale experimentation would highly benefit from a more simple and cheap way to manipulate single-cells in microfluidics systems.34 A simple microfluidic design using only a fluidic manipulation of cells has been proposed, that permits isolation of a cell in a microfabricated U-shape trap positioned in a stagnation point flow.6 The cell of interest can then be exposed to different environmental media and observed in situ. However, this system is inadequate for navigating a cell through different probing systems. In this paper, we present a very simple method to select, navigate and position a single-cell through one or a series of microfluidic probing systems.35 The building of the device requires only standard soft lithography techniques36 and a singlestep resin deposition. The technique uses a hydrodynamic control of cells, and requires no microfabricated valves, dumps or pumps. Cell motions are controlled by hydrostatic pressure using macroscopic reservoirs connected to the device and easily Lab Chip, 2010, 10, 1459–1467 | 1459
actuated on a centimetre scale. Despite extreme simplicity, the technique permits fine manipulation of cells on a micrometre scale in channels of cross section of a few tens of square micrometres (e.g. 3 10 mm), isolation of a unique cell in an analysis circuitry, self-exclusion of all undesired super-micronic particles from the analysis circuitry, and direct observation of assays with an inverted microscope coupled to a conventional 24 frames/s camera. The technique permits the minimization of the stress applied on cells, which avoids problems of mechanical damage or activation. The stress in our experiments with circulating leukocytes is, for instance, maintained at a lower level than the physiological hydrodynamic shear stress encountered in the blood circulation. Finally, the method permits observation of slow phenomena. This cheap and low-tech technique renders microfluidic single-cell manipulation and analysis straightforwardly available to a large range of applications by a wide community of biophysicists and biologists without strong expertise in microfluidics. The method for microfluidic single-cell experimentation is first described. The design rationale of the device is explained in terms of the hydrodynamic resistance of all of its constituents, and the manipulation routines for cell injection, selection and navigation in the device are described. We have demonstrated the ability of this isolation technique to integrate various microfluidic platforms by performing biomechanical and biochemical assays on human leukocytes at the single-cell level. We have exploited the method to analyze the conditions of passage or blockage of single circulating cells in the lung capillaries by mimicking the pulmonary microvasculature with 4 mm wide capillaries. We have observed the blebbing processes induced by the leukocyte deformation in the narrow capillaries. Finally, the origin of the bleb protrusions has been probed in situ by immunological staining of cytoskeleton components directly in a microfluidic device.
Fluid reservoirs were connected to the microfluidic device with polyethylene tubings (Scientific Commodities Inc., Lake Havasu City, AZ, USA) and attached to linear translation stages mounted on vertical optical rails (Micro-contr^ ole Spectraphysics S.A., Evry, France). Pressure drops across the microchannels were controlled by varying the height of the fluid macroreservoirs (Fig. 1A). Observations were taken with an inverted microscope (Olympus IX71) equipped with an oil objective of magnification 100 (Olympus, UplanFLN 1.30) and a CCD video camera (COHUE, model 4912–5000). Microfluidic sequences were recorded at 24 frames/s, digitised with a professional videotape recorder (Sony, model DSR-25) and finally
2. Experimental A. Fabrication and characterization of the microfluidic device Microfluidic channel designs were drawn using Clewin software (WieWeb Software, Hengelo, The Netherlands) to generate a chromium mask (Toppan Photomask, Corbeil Essonnes, France). The microfluidic devices were fabricated using standard soft lithography routines.36 A positive mold was created with SU-8 2015 and SU-8 2010 negative resins (Microchem, Newton, MA, USA). The resins were spin-coated on a silicon wafer (Siltronix, Archamps, France) at 3000 rpm, pre-baked at 65 C for 1 min and 95 C for 2 min, exposed at 30 mW cm2 for 10 s through the chromium mask in ‘‘hard contact’’ mode with a photolithography device (S€ uss Microtec, Munich, Germany), post-baked at 65 C for 1 min and 95 C for 2 min, and finally developed by sequential rinsing with SU8-developer and isopropanol. Ports to plug inlet and outlet reservoirs were punched with a gauge needle in the replica of the positive molds made with polydimethylsiloxane, PDMS (Sylgard 184 Silicone Elastomer Kit, Dow Corning, Midland, USA). The devices were finalized by sealing the PDMS piece on a 170 mm thick glass coverslip via O2-plasma activation (Harricks plasma) of both surfaces. All channels were incubated with a 5% Pluronic F108 solution (BASF, Mount Olive NJ, USA) for 2 h to deter cell adhesion. 1460 | Lab Chip, 2010, 10, 1459–1467
Fig. 1 Conception and modelling of the microfluidic device. (a) Schematic of the microfluidic experiment. Pressure drops are imposed by varying the height of water, DH, of the upstream (T1) or the downstream (TO) macroreservoirs, as compared to the reservoir containing the cells (T2). The inset shows a photograph of an assembled PDMS/glass microfluidic device with two entries E1 and E2, and one exit EO. The scale bar corresponds to 2 cm. (b) Micrograph of a microfluidic device with entries E1 and E2 and exit EO. Fluids injected at E1 and E2 converge at junction JC into a single channel up to junction Ji, which divides the flow between the analysis circuitry (red dashed box) and a bypass (blue dashed box). The bypass is designed to collect the largest part of the flow incoming at Ji. The scale bar corresponds to 0.33 cm. (c) Circuit analog of the device presented in (b). The two channels between entry E1 and junction JC are schematised by resistance R1, the channel between entry E2 and junction JC by resistance R2, the channel between junctions JC and Ji by resistance R3, and the bypass channel by resistance Rbp. (d) Fluid velocity in the relaxation channel as a function of the height of water in reservoir T1 (O) and TO (B). Solid lines correspond to calculations based on the circuit analog presented in (c) without adjusting parameters.
This journal is ª The Royal Society of Chemistry 2010
analyzed frame by frame using ImageJ software.37 All experiments were performed at 37 C in a home-made plexyglass box, equipped with a fan and heating resistors, enclosing the samples and the objectives. This home-made regulation ensures that the temperature of both the microfluidic device and the microscope remain equilibrated and tightly controlled. The temperature is measured in the vicinity of the microfluidic device with a thermistor and the regulation is made using Labview software. The macroscopic reservoirs used to control the hydrostatic pressure in the micro-device do not need to be in the regulated box. Each single microfluidic device was used to analyze about 50 cells. The fluid velocity in the microfluidic device has been measured by loading the device with a solution of fluorescent latex particles of diameter 0.31 mm (7-FY-300 Interfacial Dynamics Corp., Portland, OR, USA), and tracking the speed of the particles in the different parts of the device by optical microscopy. Since the velocity of the fluid depends on the distance from the channel walls, we have adjusted the focus in the centre of the channels, which permits the determination of the maximum fluid velocity in the channels. B. Reagents and cell culture The monocytic THP-1 line38 was maintained as previously described39 in Roswell Park Memorial Institute-1640 medium (Invitrogen, Cergy Pontoise, France) supplemented with 20 mM HEPES buffer, 10% Fetal Calf Serum (FCS), 2mM L-Glutamine, 10 U ml1 penicillin and 50 mg ml1 streptomycin. Stock cell cultures were passaged three times weekly and maintained at 37 C in a humidified atmosphere containing 5% CO2. The mean cell diameter was determined by cytometry at 12.5 mm with a SD of 0.8 mm. Neutrophils were extracted from whole blood samples by centrifugation in lymphocytes separation medium (Eurobio, Les ulis, France). Residual red blood cells were lysed by dilution with pure water for 30 s and debris was finally washed away. Neutrophils were then maintained at 37 C in a humidified atmosphere containing 5% CO2, and used in microfluidic experiments within 2–3 h. C. Cell fixation and labelling THP-1 cells were fixed in microchannels with a solution consisting of 4% paraformaldehyde and 0.01% Triton X-100 in PBS buffer at 37 C for 15 min and equilibrated at room temperature during incubation. Leukocytes were incubated in microchannels with anti-b-tubulin for microtubules (Sigma) and then with tetramethylrhodamine-conjugated goat anti-mouse IgG (Alexa Fluor 594, Molecular Probes). Nuclei were stained for DNA with 4,6-diamidino-2-phenylindole (DAPI) purchased from Invitrogen and actin filaments were stained with FITC-phalloidin (Alexa 488 Phalloidin, Molecular Probes, Eugene, OR). A rinsing step was applied between each staining step. Fluorescence was imaged using a Nikon Ti microscope in confocal mode or in epifluorescence mode with a Roper QuantEM 512SC EMCCD camera (Photometrics, Tucson, AZ).
3. Results and discussion Herein, we present a microfluidic technique for single-cell experimentation which is characterized by an extreme simplicity in terms of design, fabrication, and use. This technique has This journal is ª The Royal Society of Chemistry 2010
already been successfully used to study both the role of the actin network and the molecular motor activity on the passage of leukocytes in narrow constrictions.35 It has permitted the measurement of the cell deformation rate during entry in the constriction, the cell friction with the walls in the constriction and the cell shape relaxation after exit from the constriction. In the present paper, we present the technical details that should permit one to implement the method for its own investigations. The aim is to add our method to the Lab on a Chip tool box for low-throughput single-cell experiments. We first describe each manipulation step, that is the single-cell injection, navigation and selection. The principles of the method are explained based on measurements and calculations of fluid velocities in the microfluidic device. We then present new mechanical and biochemical assays of single-cell experiments using this method.
3.1 Method for single-cell manipulation a Device modelling by a circuit of hydrodynamic resistances. The microfluidic device, presented in Fig. 1a and b, has three entries: entry E1 is connected to a reservoir T1 of pure medium, entry E2 is connected to a reservoir T2 containing the suspension of cells to be analyzed, and entry EO is the output of the device connected to a reservoir TO. Fluids injected in entries E1 and E2 converge at junction JC into a single channel up to junction Ji, which divides the flow between an analysis circuitry (red dashed box in Fig. 1b) and a bypass (blue dashed box in Fig. 1b). The bypass is designed to collect the largest part of the flow ( 97%) incoming at Ji, whereas only a small portion ( 3%) of fluid enters the analysis circuitry. Although our method is adapted to any type of circuitry, we need an example of analysis circuitry to describe the hydrodynamic characteristics of the device. We use here the analysis circuitry of Fig. 1b, whose main components are a ‘‘constriction’’ channel (cross section 4 10 mm, length 2250 mm) followed by a ‘‘relaxation’’ channel (cross section 20 10 mm, length 25 mm), which has been used previously to study leukocyte trafficking through the microvasculature.35 The hydrodynamics in the device can be simply analyzed by using a lumped element modelling, as classically done with simple electrical circuits. The flow rates and pressures at each point in the device can be calculated by using an equivalent circuit of hydrodynamic resistances representative of the device, and by applying Kirchhoff’s and Ohm’s laws. Such a circuit for the device of Fig. 1b is drawn on Fig. 1c. The two channels between entry E1 and junction JC are schematised by resistance R1, the channel between entry E2 and junction JC by resistance R2, the channel between junctions JC and Ji by resistance R3, and the bypass channel by resistance RBP. The analysis circuitry itself is divided into two elements in Fig. 1c: the constriction is schematised by resistance RC and the long and wide channel by resistance RT. Each channel resistance is calculated using the solution of the flow rate in a rectangular channel (eqn (1)) with its exact dimensions, as measured with a profilometer: ! N 8HL X 1 2 W (1) Rh ¼ th bn 1mW n¼1 b4n bn H 2 where H, W and L are the height, width and length of the channel and bn ¼ (2n1)p/W. Lab Chip, 2010, 10, 1459–1467 | 1461
We define experimentally a ‘‘reference level’’ for the maximum height of liquid in the fluid reservoirs T1, T2, and TO. When the maximum heights in each reservoir are set at the ‘‘reference level’’, the velocity in all of the device should be null. Note that a fluid movement in the device in such conditions is generally the sign of the presence of a bubble in the circuitry. Bubbles greatly perturb the control of the experiment via hydrostatic pressure of water columns and must be carefully eliminated before launching an experiment. Starting with the reservoirs at the reference level, fluid movements in the device are then controlled either by lifting Ti or by lowering TO, the two other reservoirs remaining at the ‘‘reference level’’. The maximum fluid velocities measured experimentally in the centre of the relaxation channel are reported in Fig. 1d versus the height changes of reservoirs T1 and TO. These values are compared with the maximum velocities calculated in the same relaxation channel using Kirchoff’s laws at junctions JC and Ji. The perfect match on Fig. 1d between measured and calculated velocities without adjusting parameters validates the hydrodynamic description of the device. b Spontaneous filtering of micron-size particles from the analysis circuitry. The total flow Q arriving at junction Ji is divided into a major flow QBP in the bypass and a minor flow QAC in the analysis circuitry (Fig. 2 and movie 1 in the ESI†). The ratio between QBP and QAC is equal to: QBP RC þ RT ¼ QAC RBP
(2)
The device is designed to have a ratio QBP/QAC (eqn (2)) of the order of 30. The incoming channel at junction Ji is 25 mm wide. Only a small fraction of fluid, roughly 1 mm distant from the
Fig. 2 Cell selection into the analysis circuitry. (a) Micrograph of the bypass designed to collect the largest part of the flow. The red square surrounds the junction Ji, which divides the flow between the analysis circuitry and the bypass. The scale bar corresponds to 170 mm. (b) Schematic of a cell travel at junction Ji upon successive back and forth passages. The cell progressively drifts towards and eventually enters in the analysis circuitry. (c) And (d) pictures at junction Ji of the device filled with a solution of sub-micronic fluorescent beads, with observation of the zero flow condition (c) and of streamlines under flow condition (d). The scale bars for images (b–d) correspond to 17 mm.
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lower wall, enters the analysis circuitry. This explains why particles larger than approximately 1 mm, called later on ‘‘supermicronic’’ particles, do not enter the analysis circuitry on their first passage through junction Ji and are spontaneously directed toward the bypass channel. This simple and efficient technique for self-exclusion of all super-micronic particles from the analysis circuitry presents several advantages. It permits avoidance of the clogging of the analysis circuitry with too large or too numerous particles. This property is especially important when the analysis circuitry comprises narrow constrictions of a few microns width, which easily get clogged by dust particles, impurities from the cell culture medium, and debris of cells or PDMS. The self-exclusion of super-micronic particles also permits prevention of any objects or cells from entering the analysis circuitry and perturbing the single-cell experiment after the cell of interest has been injected (see movie 2 in the ESI†). On the other hand, it is interesting to note that the device does not exclude sub-micron particles. Hence, fluorescent sub-micron beads can be used to visualize the flow by optical microscopy and, therefore, measure the flow velocity in all channels of the microfluidic device (Fig. 2a–c and movie 1 in the ESI†). Note that the flow focusing geometry at junction JC is also designed to help the filtering efficiency at Ji. The fluid incoming from E1 is first filtered at 2 mm before injection in the microfluidic device and then filtered again at 10 mm in the microfluidic device. Consequently, the fluid coming from E1 is free of cells and other particles larger than 2 mm. The main source of particles susceptible to clog the analysis circuitry is, therefore, coming from E2. Indeed, the suspensions loaded in E2 cannot be filtered because they contain the cells. In this context, the flow focusing at Ji tends to centre the particles incoming from E2 in the channel between JC and Ji. This central position of particles increases the probability for particles to be directed toward the bypass at junction Ji. Conversely, an injection forcing the cells toward the lower wall would increase the probability for cells to enter the analysis circuitry. In practice, we observe with the device in Fig. 1b that 100% of cells and particles larger that 1 mm are spontaneously directed toward the bypass channel on their first passage at Ji. c Injection of cells in the microfluidics device. Injection of cells in a microfluidic device is not straightforward due to the fact that their sedimentation velocity is generally larger than the fluid velocity in the macroscopic circuitry. To give an order of magnitude, the total flow rate in the device, when reservoir T1 is lifted 10 cm above reference level and the two other reservoirs are positioned at the reference level, is of 5.041014 m3 s1 (i.e. 4.7 mL per day). As a consequence, cells are generally not driven into the microfluidic device by the fluid motion. We use here a simple procedure for cell injection in the microfluidic device, which takes advantage of the preponderance of the cell sedimentation. We start with the microchannels fully loaded with cell culture medium, and the three reservoirs positioned at the ‘‘reference level’’ in order to stop the flow in the system (Fig. 2c). The reservoir T2 and tubing of port E2 are then unplugged from the device and filled with a cell suspension at 106 cells mL1 by suction. The suction procedure ensures that the cell suspension is present at the extremity of the injection tubing. The reservoir T2 and the tubing are then re-plugged gently into port E2 by taking This journal is ª The Royal Society of Chemistry 2010
care that no bubble is trapped during the process and that the tubing is maintained vertically. The three reservoirs being level, the flow is negligible and the cells from the injection tubing sediment at the bottom of port E2. The first cells touch the bottom within a few minutes. In order to inject cells in the device, the outlet reservoir TO is then lowered by a few tens of centimetres. In these conditions, the flow velocity in the vicinity of the microfluidic channel entrance of port E2 is high enough to carry cells lying at the bottom of port E2. A cell of interest is then navigated in the microfluidic channels by manual actuation of reservoir TO on a few centimetres scale. The cell motion is followed by direct observation with an inverted microscope from their entry at port E2 up to their arrival at junction Ji. All cells that are sucked in the microfluidic at E2 and that are not selected for an experiment will eventually travel in the bypass channel to the output reservoir TO, and will, therefore, not clog the analysis circuitry. d Cell selection into the analysis circuitry. On its first passage at junction Ji (Fig. 2b), a cell is spontaneously directed into the bypass channel. In order to force the entry of a cell into the analysis circuitry, one has to simply navigate the cell back and forth in the vicinity of the corner of junction Ji by alternatively moving the reservoir TO up and down. Upon a few successive passages, the cell progressively drifts towards the analysis circuitry channel and finally enters (Fig. 2b and movie 2 in the ESI†). According to the properties of Stokes flow, all fluid motions are expected to be reversible. In the absence of cell at junction Ji, fluid particles always follow the same streamline, either in the bypass or in the analysis circuitry, when the flow is driven back and forth. On the other hand, when a cell is present in the flow, its surface is involved in the flow boundary conditions and the travel of the cell is not reversible upon successive passages. The calculation of a cell path in a complex geometry with moving boundaries is a difficult task. Qualitatively, upon successive passages, the cell is progressively sucked in the analysis circuitry. The time required to inject a cell in the device and isolate it in the analysis circuitry takes only about 10–30 s. e Precise cell navigation in analysis circuitry. The device permits an easy and precise navigation of cells in the analysis circuitry by manual displacement of reservoirs on a centimetres scale. The manipulation is fine enough to navigate cells in narrow environments, such as a constriction channel of cross section 3 10 mm (e.g. movie 3 in the ESI†). The cell can be stopped in the constriction with negligible drift (< 1 mm min1). The possibility to slow down the travel in the micro-circuitry permits one to record experiments with conventional 24 frames/s camera, which presents several advantages. Indeed, there is no need for an expensive fast acquisition camera. More importantly, all phenomena can be observed live. This allows, for instance, the opportunity to select a cell of interest in a wide population. Working at low fluid velocity is also important to limit the hydrodynamic shear stress on cells during their navigation in the microchannels. Indeed, shear stress is known to be a cause of activation49,54 and damage of circulating cells. f Fine and quantitative control of fluid velocities and pressures. The precision of the control of very low flow velocity in the analysis circuitry results from the precise hydrostatic pressure This journal is ª The Royal Society of Chemistry 2010
control, and also from the important flow reduction in the analysis circuitry at junction Ji. For a given hydrostatic pressure imposed with the macroscopic reservoirs, the flow rate in the analysis circuitry is 30 times smaller than in the rest of the device. On top of that, our device allows two different levels of control, either fine or coarse, by actuating either the reservoir T1 or the reservoir TO. Lowering TO from the reference level position imposes fluid velocities 5 times larger than lifting T1 by the same height. The flow calculation based on the equivalent resistive circuit fits perfectly this experimental behaviour (Fig. 1d). The pressure drop Pi PO between the inlet and the outlet of the 4 mm wide constriction of Fig. 1c can be expressed against the pressure of the reservoirs as: ! RC R3 aP3 þbR2 P1 þbR1 P2 Pi - Po ¼ - P3 (3) RC þ RT 1 þ aR3 -bR1 R2 where a¼
1 1 1 þ and b ¼ RC þ RT RBP R1 R2 þ R1 R3 þ R2 R3
(4)
The calculation shows that Pi Po is 10 times smaller than the pressure drop P1 imposed by lifting T1 from the position ‘‘reference level’’. One centimetre actuation of T1 applies an effective pressure drop of 10 Pa across the constriction. This precision range is well adapted to mimic physiological pressure drops encountered in the human microcirculation. If required in other applications, higher precision can be obtained by designing a device with a set of hydrodynamic resistances providing a higher pressure drop demultiplication. One has to underline that the pressure across the microfluidic constriction is not only precisely controlled via actuation of the height of the macroscopic reservoir, but it is also independent of the presence of cells in the microfluidic constriction. Indeed, the blockage of the flow by the arrest of a cell in the analysis circuitry (e.g. a constriction) perturbs the flow rates and the pressure gradients in the circuitry. However, one has to remember that the analysis circuitry has a much higher hydrodynamic resistance than the bypass. The change from a very resistive to an infinitely resistive (i.e. clogged) analysis circuitry does not noticeably perturb the flow rate in the bypass: the change of pressure drop between Ji and EO is negligible, around 3%. This sheds light on another advantage of the bypass geometry: it ensures that the pressure drop across the analysis circuitry during an experiment is only dependant on the pressure applied externally by the macroscopic reservoirs and not on the presence of cells or other objects in the circuitry. g Acquisition speed. The main goal of our method is to achieve a cheap and user-friendly single-cell experiment device that can be entirely actuated manually by macroscopic displacements of reservoir heights on a scale of centimetres. The method is, therefore, not designed to perform automated high throughput measurements. Nevertheless, several tens of cells can be analyzed per hour with the present device, which allows enough statistics for many lab-scale research experiments. On the other hand, the ability to work at low velocity is also an advantage in certain experiments, e.g., for the study of slow phenomena such as activation events, membrane unwrinkling, bleb formation, or shape Lab Chip, 2010, 10, 1459–1467 | 1463
relaxation. This method is of special interest for microfluidic tools designed to study cell mechanics40 or immunostaining of cells of interest directly in a microfluidic device. 3.2 Mechanical and biochemical assays on single circulating cells Recent studies on neutrophil chemotaxis41 or tumour cell deformation and migration38,42 have confirmed the interest of microfluidics for quantitative cell biology. Despite these successful experiments, the use of single-cell microfluidic assays is still limited, mainly due to the difficulties in isolating circulating cells. A key issue for the use of complex microfluidic geometries relies on the efficiency of the cell isolation method to avoid capillary clogging, cell–cell interactions and modification of hydrodynamics conditions during experiment. We show here how our microfluidic method can be useful to perform microrofluidic mechanical and biochemical experiments on single circulating leukocytes in constriction channels. Former studies have raised questions regarding the precise role of mechanical properties and activation processes of white blood cells during their passage in the microvasculature. In particular, the mechanical plugging of lung microvessels by leukocytes has been implicated in the physiopathology of several diseases, such as the Acute Respiratory Distress Syndrome (ARDS), which results in a mortality rate around 50%.43 The direct visualisation of leukocyte retention characteristics in human microvasculature is, in general, very difficult, except with a limited resolution in tissues like the retina44,45 and the tongue,46,47 and is altogether impossible in the pulmonary microvasculature. In this context, mimetic in vitro assays using microfluidics is a new and very helpful approach, which has recently brought new insight into these problems.4,35,42,48–52 The in vitro approach also presents specific advantages, such as the possibility to control experimental conditions precisely (flow, pressure, cell characteristics, medium,.) or to test new drugs before testing on humans.
a Single-cell mechanical behaviour in narrow capillaries. We first present an experiment with the single-cell analysis circuitry of Fig. 1c, i.e., a long constriction of width 4 mm. This device permits observation of the cell deformation rate during the entrance stage, the cell velocity during the transit stage in the constriction, and the shape recovery during the relaxation stage after cell release from the constriction.35 We have made experiments using this device with primary neutrophil cells, which are very sensitive to chemical or mechanical stresses54 and are known to get easily activated.53 These experiments show that our method of cell injection, navigation and selection permits avoidance of mechanically induced damage or blebbing of cells, even with neutrophils (movie 3 in the ESI†). Further details on results obtained with this device have been presented elsewhere.35 We present here new results obtained with another analysis circuitry made of a succession of 50 interconnected capillary segments of 4 mm width connected to 20 mm wide inlet and outlet channels. This device permits the study of the dynamics of single leukocytes through interconnected capillary segments (Fig. 3). The time required for passage through the capillary network of a single alveolus could be assessed for each leukocyte, and represents its alveolar capillary transit time. Our results show that alveolar capillary transit times range over two orders of magnitude under the same pressure drop (see Fig. 3a and 3b). Leukocytes can either pass through the capillary segments in a very short time (Fig. 3b, t ¼ 120 ms) or stop for long periods of time (Fig. 3a, t ¼ 12 500 ms). Our observations indicate that the temporary stops are affected by the shape of the leukocyte and the initial orientation of cells at capillary entry. Indeed, the leukocytes have a squeezed shape at the exit of a capillary and resemble a puck. If a puck-shape cell presents its smallest dimension to the entrance of the next capillary, the time required to pass through this capillary segment is very short (see Fig. 3b). Conversely, if the puck-shape cell presents its largest dimension at the entrance of the next capillary, the cell is temporarily stopped (movie 4 in the ESI†). In the latter case, the cell has to
Fig. 3 Mechanical investigation. (a) If the puck-shaped cell presents its largest dimension to the entrance of the capillary, the time required to pass through a capillary segment is long (t ¼ 12 500 ms). This situation corresponds to a temporary stop. (b) If the puck-shaped cell presents its smallest dimension to the entrance of interconnected capillary segments, the time required to pass through a capillary segment is very short (t ¼ 120 ms). (c) Experiment perturbed by the entry in the device of a doublet of cells (see also movie 5 in the ESI†). The scale bar corresponds to 8 mm.
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deform and adapt its shape to the capillary entry. The entrance time after such a stop is then comparable to the cell entrance time in the first capillary. Perturbations of cell orientation, coupled to the slow process of cell deformation, can, therefore, result in important retention times of cells in the multiple constrictions of the microvasculature. However, the sequential constriction of the real microvasculature, e.g. in the lungs, does not have the sharp crenellations of our microfluidic device. A smooth constriction entrance will help a deformed cell to present its smallest dimension toward the entrance. Sequential stops in sequential constrictions is, therefore, not likely. This suggests that the slow shape recovery (around 30 s) will rather favour a fast in vivo passage of cells through successive capillary segments. We want to underline here that our method is not limited to experiments where flow is actually required for the analysis of the isolated cell, such as in the mechanical experiment described here. Selected cells can also be manipulated by repeatedly reversing flow to iteratively measure a property of interest with different tools. Selected cells can also be trapped in the device and exposed to different environmental media. This latter possibility permits one to fix and stain cells in situ as demonstrated below (paragraph c). b Importance of cell isolation. The importance of the spontaneous isolation of the analysis circuitry from all undesired super-micronic particles is illustrated on Fig. 3c (see also movie 5 in the ESI†). This experiment was performed with a device having a bypass that is not optimised to prevent the entry of super-micronic particles in the analysis circuitry; which permits visualization of certain problems that our method permits to avoid. Fig. 3c (and movie 5 in the ESI†) shows the passage of a doublet of leukocytes in strong mutual adhesion. One can see that the transit of a doublet of cells is very complex and also noticeably slower than the transit of a single cell. The passage of aggregates of several cells is even more difficult and results generally in an irreversible clogging of the device. Obviously, cell
selection with our method permits avoidance of the entry of such aggregates, which are not rare in leukocytes suspensions. It is known that mechanical stimulation can promote cellular activation and, therefore, increase the tendency of cell adherence. The channels’ and tubings’ surface has been treated with Pluronic F108 to deter cell adherence to the circuitry walls but aggregation of cells with eachother cannot be totally avoided. It is, therefore, necessary to isolate by default the analysis circuitry from any large circulating objects. Another advantage of isolating the analysis circuitry from circulating cells and objects is to prevent the entrance of undesired objects in the analysis circuitry during an experiment with a single cell. This is illustrated at the end of movie 5 in the ESI,† where the arrival of a third cell in the channel during a experiment with a doublet of cells perturbs noticeably the passage characteristics of the doublet itself. c Biochemical investigation of blebbing. A large number of studies have suggested that activation of leukocytes could be involved in their capillary sequestration.54,55 Conversely, mechanical deformation of leukocytes in capillaries can be responsible of their activation and trigger the formation of pseudopods or blebs. Although less studied than lamellipodia or filipodial protrusion, blebbing is a common activation phenomenon encountered during apoptosis, cytokinesis, cell motility or cell spreading. The origin and the specific role of blebbing in physiological and pathological situations is still subject to debate. Most mechanistic knowledge on bleb formation has been built on adherent cells. However, the study of the blebbing process of circulating cells may also be essential to understand the flow behaviour of tumour cells or leukocytes. The contribution of the activation process on cell mechanical properties has been extensively studied using chemotactic agents, and different techniques like filtration, pore transit or microindentation. Yap and Kamm have recently shown using pore transit experiments that a combination of low driving pressures
Fig. 4 Bleb formation during the shape recovery stage. Image sequence showing the formation of membrane protrusions (blebs) following the deformation of a THP-1 cell through a 4 mm wide channel. White arrowheads point to the growing blebs. The scale bar corresponds to 10 mm.
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and small capillary diameters can promote cellular activation, or more precisely blebbing, of circulating leukocytes.54 However, this technique could not produce a direct observation of the blebbing process of circulating leukocytes. The study of activation processes in physiological flows requires a microfluidic manipulation technique that is gentle enough to maintain hydrodynamic shear stress at a level that is low enough to avoid the damage and/or activation of cells. This is made possible with our method, which allows navigation of cells at very low velocities. The method has, for instance, permitted us to observe the whole blebbing process of single leukocytes after their transit through a narrow microfluidic capillary of width 4 mm and length 2250 mm. The results are presented on Fig. 4. The blebbing process consists of the growth of roughly spherical protrusions on the cell surface, which occur mostly after release from the constriction during the shape relaxation stage (Fig. 4e–l). The expansion of blebs leads to a large deformation of the leukocytes (Fig. 4l). Then, membrane protrusions stop growing, retract (Fig. 4m–o) and eventually disappear (Fig. 4o). The transition from bleb expansion to stasis and finally contraction occurs over 30–50 s. Hence, blebs are dynamic and transient but their formation and contraction coordination remains an open question. A possible explanation comes from recent studies,56,57 which propose that filamentous actin (F-actin) contributes to bleb formation. Immunocytochemical imaging is the method of choice to shed light on the role of actin filaments. Our microfluidic method permits performance of immunocytochemical staining directly in the microdevice on the circulating cell of interest after the mechanical assay. As an example, we have stained the different cytoskeletal components (actin filaments, microtubules and nucleus) in normal (Fig. 5a) and blebbed leukocytes (Fig. 5b). Images of deformed leukocytes show clearly the formation of membrane protrusions (see white arrows on Fig. 5b). Therefore, the high level of F-actin in blebs indicates that filamentous actin is involved in the formation of blebs, which validates the role of actin in bleb expansion. On the other hand, it is interesting to note the absence of microtubules in
Fig. 5 Immunostaining investigation by confocal microscopy. Confocal images in the microfluidic device of (a) a normal leukocyte and (b) a deformed leukocyte. Both are stained in suspension for F-actin in green, nucleus in blue and microtubules in red. White arrows on (b) show a high level of F-actin in bleb protrusions which confirms that F-actin is involved in the formation of blebs, while the absence of microtubules in membrane protrusions suggests that they are not involved in the formation of blebs. The scale bars correspond to 8 mm.
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membrane protrusions (Fig. 5b), which suggests that microtubules are not involved in the formation of blebs of circulating leukocytes. Membrane detachment and bleb inflation are, therefore, thought to be driven by intracellular pressure transients generated by the actin cortex. How precisely this phenomenon is initiated by the myosin II activity deserves to be resolved in the future by using this isolation technique, for instance with drugs inhibiting actin organization or myosin II activity. Our microfluidic method provides a well adapted tool to perform these systematic investigations.
4. Conclusion and outlook Single-cell experiments permit one to go beyond cell population averaged measurements and get access to dynamic cell phenotypes at cellular and sub-cellular level. Microfluidics is a method of choice for single-cell experimentation. However, existing microfluidics methods for single cell experimentation involve often sophisticated devices that permit full automation of complex tasks, but somehow hampers the wide spreading of microfluidic single-cell experimentation to many research labscale assays. Our very simple and cheap method permits manipulation of single-cells in microfluidics systems and allows non-specialists in microfluidics to easily design their own singlecell experiments. The method is based on a flow division between the single cell analysis circuitry and a bypass, which permits one to maintain the analysis circuitry clear of all supermicronic particles, to isolate a cell of interest in the analysis circuitry, to navigate the cell with precision in narrow channels by manual actuation of macroscopic fluid reservoirs, and to observe cellular phenotypes in microfluidic devices on a slow timescale with a conventional 24 frames per second camera. The cell selection and navigation routines of the method induce no perturbation susceptible to damage or activate sensitive cells like neutrophils. Several tens of cells can be analyzed per hour, which yields satisfactory statistics for many lab-scale research experiments. The method is particularly adapted for experiments in channels with dimensions of a few microns and for the study of slow phenomena, such as activation, friction in a constriction, membrane unwrinkling, bleb formation, or shape relaxation. It was used here to manipulate single leukocytes in a series of constrictions to investigate circulating cells’ properties under the mechanical constraints encountered in the microcirculation. This approach has yielded direct insight into the influence on the transit times of cell orientation at the entries of successive constrictions, and into the cell blebbing process induced by mechanical stress. Immunostaining of cells directly in the microfluidic device is also enabled by our method. It has permitted us to demonstrate that F-actin microfilaments, and not microtubules, are present in the blebs of circulating cells. This suggests that blebbing is initiated by the same process on adherent and circulating cells. Although we have focused on applications of our method to cell mechanics and immunostaining assays, we expect that our method will be of interest for other single-cell assays probing other cell phenotypes, and also for experiments with other micron-scale objects than cells, e.g. vesicles, drops, or aggregates of bacteria and cells. This journal is ª The Royal Society of Chemistry 2010
Acknowledgements The authors gratefully acknowledge the Rhodia Company (LOF, Pessac, France) for financial support of SG’s CNRS postdoctoral position. We also thank Mathieu Guirardel and Pascal Silberzan for technical support and advice with microfabication issues. This work was partly supported by the National Fund for Scientific Research (F.R.S.-FNRS) through ‘‘MIS Confocal Microscopy’’ and ‘‘Credit aux Chercheurs’’ grants, and by a grant from the French C-NANO network in Region PACA. Doctoral fellowships are supported by F.R.I.A. for MV, and by the Region PACA and the company CAPSUM-SAS for PP.
Notes and references 1 2 3 4
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