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Sub-cellular precision on-chip small-animal immobilization, multi-photonimaging and femtosecond-laser manipulation†

Fei Zeng,‡ Christopher B. Rohde‡ and Mehmet Fatih Yanik*

Received 20th March 2008, Accepted 28th March 2008First published as an Advance Article on the web 2nd April 2008DOI: 10.1039/b804808h

Techniques for stable, rapid and repeatable small-animalimmobilization are necessary for high-throughput in vivogenetic/drug screens using cellular and sub-cellular featuresin multi-cellular organisms. We demonstrate a method fornon-invasive and high-throughput on-chip immobilizationof physiologically active C. elegans without the use ofanesthesia or cooling, but with comparable stability evenfor the most demanding purposes. We show observationand manipulation of sub-cellular features in immobilizedanimals using two-photon microscopy and femtosecond-laser microsurgery.

C. elegans, the small semi-transparent nematode, is a powerfulmodel for studying a wide variety of biological phenomena. Ithas significantly impacted many areas of cellular biology, whichis exemplified by the two recent Nobel prizes awarded within thelast six years for the discoveries made using this organism. Cur-rently, C. elegans is being used to study many disease models andmulti-cellular processes including Parkinson’s and Alzheimer’sdisease, muscular dystrophy, aging, fat metabolism, and innateimmunity. C. elegans′ small size (∼50 lm diameter, ∼1 mmlength), transparency, amenable genetics, hermaphroditic andgonochoric reproduction, and rapid developmental cycle all con-tribute to this. However, the high mobility of animals requirestheir immobilization for imaging and manipulation of cellularand sub-cellular features necessary for study of most biologicalprocesses and disease models. Immobilization is most commonlydone using anesthesia. Anesthetics affecting C. elegans includesodium azide (NaN3), levamisole and tricaine/tetrimisole. Un-fortunately, the side-effects of anesthesia on many biologicalprocesses are detrimental or uncharacterized. For example, theability of C. elegans to survive anesthesia by NaN3 has beenlinked to the induction of the heat shock proteins Hsp70 andHsp16 and increased thermotolerence.1 Significant cooling ofanimals can also immobilize them reversibly,2 but the effectsof cooling on the biological processes are also similarly unpre-dictable, and many worm strains show sensitivity to temperaturechanges. Furthermore, anesthesia is not suitable for high-throughput screening3,4 and the lack of techniques for rapid andrepeatable small-animal immobilization is significantly limiting

Massachusetts Institute of Technology, Department of ElectricalEngineering and Computer Science, Cambridge, MA, 02139, USA.E-mail: yanik@mit.edu† Electronic supplementary information (ESI) available: Device fab-rication, cell-body tracking, lifespan synchronization, animal prepa-ration, and lifespan analysis and supplementary movie. See DOI:10.1039/b804808h‡ Equally contributing authors.

high-throughput in vivo genetic/drug screens involving cellularand sub-cellular features.3,4 Anesthesia is also not suitablefor studies requiring physiologically active animals, such asthose examining neurophysiology, germ-line proliferation ordevelopment. Thus, a method of immobilizing physiologicallyactive animals precisely, repeatably and rapidly with minimalphysiological effects is of great importance.

Many of the properties that make C. elegans a useful modelorganism also make it amenable to manipulation in microfluidicchannels. A number of novel microfluidic devices to studyC. elegans have been published, including mazes for studyinglearning,5 devices for the generation of oxygen gradients,6

optofluidic imaging platforms,7 automated cultivation systems,8

and a shadow imaging system for studying animals in space.9

Recently, methods for partially immobilizing C. elegans usingmicrofluidic devices have been reported. These include theuse of an array of small aspiration channels to immobilizeanimals in a high-throughput platform,10 and tapered channelsfor immobilizing either single11 or multiple12 animals. Bothtechniques can be used to immobilize animals to a degree wherethe movement is small for brief periods. However, this is notsufficient for highly reliable imaging and manipulation of cellularor sub-cellular features using techniques such as high-resolutiontime-lapse microscopy, multi-photon and confocal imaging, andfemtosecond-laser microsurgery.

Here, we present a technique for rapid, repeatable andextremely stable immobilization of physiologically active ani-mals without anesthesia or cooling that allows imaging andmanipulation of sub-cellular features for the most demandingpurposes. Our technique achieves stability comparable to that ofdeep anesthesia and affects neither the lifespan of the animalsnor their brood size, and does not induce hypoxia. Our methodcan immobilize animals in fractions of a second, and can bereadily integrated with the microfluidic systems we previouslydemonstrated for high-throughput small-animal screening,10

where animals can be temporarily immobilized at different time-points for imaging and manipulation.

Our devices consist of multiple thin layers of poly(dimethylsiloxane) (PDMS) fabricated by soft lithography13 (see the ESI†).A 100 lm-tall ‘flow’ channel contains multiple 15 lm-tallaspiration channels that capture/align the animals in a linearposition when the pressure in the aspiration channels is lowered(steps i→ii in Fig. 1(a)). The aspiration immobilizes animalsonly partially, and it is not sufficient to completely restrict theirmotion. In order to fully immobilize the animals, we create aseal around them that restricts their motion completely. Thisis done by using a 15–25 lm-thick flexible ‘sealing’ membranethat separates a ‘press-down’ channel from the ‘flow’ channel

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Fig. 1 Microfluidic immobilization of C. elegans. (a) Illustration ofthe immobilization process. (i) Without suction, the worm is free tomove in the microfluidic channel. (ii) The pressure is lowered in themultiple aspiration ports on the left side of the ‘flow’ channel to partiallyimmobilize the animal against the left side of the ‘flow’ channel. (iii)The fluidic pressure in the ‘press-down’ channel is increased, flexingthe membrane separating the channels downwards, sealing the animalagainst the multiple aspiration ports and fully immobilizing it. (b)Low magnification image of a worm immobilized in the microfluidicdevice, showing the multiple suction ports on the left side of the flowchannel (scale bar 250 lm). (c) Close up of immobilized pmec-4::gfp an-imal, showing gfp-labeled fluorescent posterior lateral mechanosensory(PLM) neurons (scale bar 20 lm).

underneath. The press-down channel can be rapidly pressurizedto expand the thin membrane downwards, as in microfluidicvalves.14 The membrane flexes on top of the captured animals,wrapping around them and forming a tight seal which com-pletely constrains their motion in a linear orientation. Althoughthe animals are constrained by the PDMS membrane from thetop and bottom, they still have access to liquid media via themultiple aspiration channels on the side. Fig. 1(b) shows animage of an immobilized adult animal in the device, and Fig. 1(c)shows superimposed bright-field and fluorescence images takenat high magnification. A movie of the full immobilizationprocess, with the duration of each step increased for ease ofvisualization, is available in the ESI.† The movie shows a singleaspiration port across from the multiple-aspiration ports, whichallows capture of one and only one animal, and subsequentwashing of the channels before the multiple-aspiration channelsare turned on.10 This prevents simultaneous immobilization ofmore than one animal. A ‘control’ layer below the flow layer(not shown in Fig. 1) allows us to switch pressures on variousports rapidly via push-up microfluidic valves.14

We have performed several qualitative and quantitative as-sessments of the degree of immobilization. Fig. 2(a) illustratesan animal’s motion at three different time points followingimmobilization. Images of the anterior ventral mechanosensory(AVM) cell body and its axon are taken 5 s apart, pseudo-coloredred (first time point), green and blue (final time point), andthen superimposed. White regions indicate areas overlappingin all three time points. As seen in Fig. 2(a), the degree ofmovement for an animal immobilized in our device is small,even at 50× magnification. To quantify the effectiveness ofour microfluidic immobilization versus regular anesthesia, wetracked the cell bodies of touch neurons labeled with greenfluorescent protein (GFP) using a software algorithm (see ESI).†C. elegans has six touch neuron receptors, with three cellsdetecting anterior touch, located in the anterior mid-body, and

Fig. 2 Quantitative and qualitative measurements of microfluidicimmobilization. (a) Positional stability of a sub-cellular feature duringmicrofluidic immobilization. Three images of the anterior ventralmechnosensory (AVM) neuron, its axon, and the PVM axon are takenat 5 s apart. These images are then colored red (earliest), green, and blue(latest) and overlaid on top of each other, such that white regions indicatethe overlap between all three images (scale bar 20 lm). Inset shows close-up of outlined area. (b)(c) Comparison of cell body movement duringmicrofluidic immobilization versus deep anesthetic immobilization using10 mM NaN3. The centroids of the touch neuron cell bodies aretracked using a computer algorithm (see ESI)† that analyzes data frommovies taken at 20–30 Hz with 50× magnification. 12 microfluidically-immobilized animals and 9 anesthetized animals were tracked. (b)Histogram showing average displacement of cell bodies between framesdivided by the time between frames. (c) Line plot showing mean drift ofcell bodies over time.

three cells detecting posterior touch, one located in the posteriorventral mid-body and two in the tail. The results of tracking thesecell bodies are shown in Fig. 2(b), which shows the histogram ofthe frame-to-frame displacement of the tracked centroids, andFig. 2(c), which shows their mean drift over time. These resultsshow that the movement of the immobilized animals is quitesmall, and is comparable to their motion even when deeplyanesthetized. Despite being completely restrained externally,the animals can still have small internal movements, especiallyaround the pharynx. However, such internal activity does notcause significant displacement of the neurons imaged, and wewere able to acquire high-resolution two-photon images evennear the pharynx, as shown below.

To check whether our device injures the animals duringimmobilization, we tracked the lifespan and brood size of 25animals that were each immobilized for 1 min using 15 PSI ofpressure in the ‘press-down’ channel (see ESI).† The pressureapplied to the animals by the membrane is lower than this, dueto the resistance of the membrane to flexing. The immobilizedpopulation was compared to a control population that was notrun through the device. The lifespans of both populations at20 ◦C are shown in Fig. 3. The mean lifespan of the immobilizedpopulation was 17.3 days (s.d. =.05 days) and 16.9 days for thecontrol population (s.d. = 4.0 days). We used the GraphpadPrism software package to perform the log-rank (Mantel-Cox)test, which gave a p-value of 0.8947. This suggests there isno statistically significant difference between the lifespans ofthe two populations. Both populations also produced normalbrood sizes, and the immobilized population was free of axonal

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Fig. 3 Lifespans of microfluidically-immobilized and control popu-lations. The immobilized population consisted of 25 worms that wereimmobilized for one minute each, and the control population consistedof 23 worms that were not run through the device. Both populationswere monitored once a day for dead animals, and surviving animalswere transferred to a fresh plate. An animal was scored as dead if it didnot respond to prodding with a platinum worm pick (see ESI).† Themean lifespan of the immobilized population was 17.3 days (s.d. 5.0days), and the mean lifespan of the control population was 16.9 days(s.d. 4.0 days).

blebbing, suggesting that the animals did not become hypoxicwhile immobilized.15

Our method allows highly demanding sub-cellular resolutionimaging and manipulation techniques requiring long periods ofstable immobilization. We have chosen two applications thatdemonstrate this: femtosecond-laser microsurgery and three-dimensional two-photon microscopy.

Femtosecond-laser micro/nanosurgery enables precisionablation of sub-cellular processes with minimal collateraldamage.16 We have previously employed this technique to per-form the first axonal regeneration study in C. elegans,17,18 whichopened up tremendous potential for genetic/drug discoverieson neural degeneration and regeneration using a geneticallyamenable whole-animal model. However, manually preparingan animal for surgery, imaging and recovering it afterwards arefairly laborious tasks, and the ability to rapidly immobilize ani-mals in a repeatable fashion can greatly accelerate investigationsinto neural degeneration and regeneration. Furthermore, theeffects of long-term anesthesia on these processes are not known.Using our immobilization technique, we can immobilize animalsand perform femtosecond-laser microsurgery repeatably, rapidlyand with sub-cellular precision. Fig. 4(a) shows an image ofa pmec4::gfp animal whose touch neuron process has beencut. Our on-chip femtosecond-laser microsurgery technique is apowerful tool for the discovery of potential drugs and geneticfactors affecting neural degeneration and regeneration.

Multi-photon microscopy, including two-photon excitationfluorescence (TPEF),19 is another important application that re-quires a high degree of stabilization. Multi-photon microscopy isinherently non-linear, and thus has the ability to perform opticalsectioning with negligible out-of-plane absorption and emission.This dramatically reduces photobleaching and phototoxicity.20

This is especially significant in assays that require animals to beimaged at multiple time points. The advantages of multi-photonmicroscopy are offset by the requirement for live animals tobe anesthetized, due to the long duration needed to capture animage. We have used our immobilization method to successfullyacquire two-photon images of non-anesthetized live animalsusing devices bonded to 175 lm thin glass slide. Fig. 4(b)and Fig. 4(c) show two volume reconstructions of pmec4::gfp

Fig. 4 Microfluidic immobilization enables several key applications inlive, un-anesthetized animals. (a) Successful axotomy of AVM process.20 nJ femtosecond pulses were delivered using a Mai Tai R© Ti:Sapphirelaser (Spectra-Physics) at a rate of 80 MHz to successfully cut the axons.Arrow is indicating the focus of the laser and the axotomized region.(b), (c) Volume reconstruction of images captured using two-photonmicroscopy. (b) Head of mec4::gfp animal. (c) Posterior midbody ofmec4::gfp animal and posterior ventral mechanosensory neuron (PVM).The left and right anterior lateral mechanosensory (ALML and ALMR)and AVM processes are clearly visible, as are the nerve ring branchesthat extend into multiple imaging planes (all scale bars 20 lm).

animals obtained by a two-photon microscope scanning at 0.2frames s−1 using a 40×/0.8 NA water-immersion objective. Inthis configuration, imaging a 120 lm × 120 lm × 30 lm volumerequired roughly two minutes of stable immobilization. Suchnon-invasive imaging of non-anesthetized animals can allowinvestigation of cellular processes sensitive to anesthesia, suchas neural degeneration, regeneration and embryogenesis.

Here, we demonstrated a rapid and highly repeatable tech-nique for immobilizing small animals for imaging and manipu-lation of sub-cellular processes without anesthesia. The stabilityof our technique is comparable to that of deep anesthesia.It affects neither the lifespan nor brood size of the animals,and it does not induce hypoxia. Our on-chip method canenable high-throughput screening of cellular and sub-cellularphenotypes in whole animals, as well as the use of high precisiontechniques such as femtosecond-laser microsurgery and multi-photon microscopy on physiologically active animals.

Acknowledgements

This work was supported by the NIH Director’s New Innova-tor Award Program (1-DP2-OD002989–01), part of the NIHRoadmap for Medical Research. We thank P. So and J. Cha(Massachusetts Institute of Technology) for assistance with two-photon imaging.

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