3D manipulation of small multicellular organisms in soft microtubes

Caenorhabditis elegans (C. elegans) and Danio rerio (zebrafish) are established as essential model organisms for biological assays to model human diseases and screen new drugs. However, their small size poses a challenge when it comes to adjusting their body orientation for imaging¹ or microinjection.² The most common approach currently is manual handling with tweezers or a fine tip under a stereoscope.³ This method is labor-intensive and difficult, prompting the development of microfluidic devices for manipulation based on their comparable length scales.

Microfluidic devices can immobilize these organisms using pneumatically actuated flexible PDMS membranes⁴ or trapping them in narrow constriction channels.⁵ However, controlling their body orientation within a microchannel remains challenging and requires sophisticated passive and active techniques. One passive technique for C. elegans involves integrating a curved channel with a radius of curvature of 125 μm, achieving laterally oriented animals with 84% efficiency.⁶ An active method uses microfluidic vortices to rotate the worm, generating stepwise body rotation through a controlled pulsatile flow.⁷ However, this method requires prior paralysis of the worms with levamisole to ensure a straight body shape. Another active method uses acoustic waves to control the orientation of C. elegans.^8,9 A hybrid microfluidic device incorporating a rotatable glass capillary has been developed for flexible orientation control. The head or tail is captured with the glass capillary, rotated manually, and immobilized for imaging using side suction channels.¹⁰ 

For zebrafish larvae, a method involving trapping embryos in a fluorinated ethylene propylene (FEP) tube with low concentrations of agarose (0.1%) or methylcellulose (3%) has been proposed.¹¹ However, such immobilization can impact growth, making agarose-free methods more desirable. The Zebrafish Entrapment by Restriction Array (ZEBRA) device fine-tuned channel geometry to achieve positioning without agarose.¹² Another device, the “Fish-Trap” array, traps a single larva’s tail in a restricted portion, allowing body orientation control between lateral and dorsal without anesthetics.¹³ However, stepwise positioning was not demonstrated.

To address these challenges, we propose a new technique using a transparent, flexible microtube under mechanical stretching to manipulate and immobilize organisms. This method allows for temporary immobilization and rotation of the organism for comprehensive imaging. After imaging, the organism can be released and collected for further analysis. The soft microtube is easy to fabricate and use, even by non-trained users, due to its similarity to traditional glass capillaries. This soft microtube could become a versatile tool for laboratories working with small multicellular organisms.

To build transparent and elastic tubes for C. elegans (∼50 μm in diameter), we adapted an electrothermal curing method from previous work.¹⁴ We used a mixture of 30 g PDMS and 3 g curing agent (10:1 ratio), which was degassed for 90 min. A 20 cm-long nichrome resistance wire, ∼78 μm in diameter, served as the core of the hollow tube. A 1.5 cm-long polytetrafluoroethylene (PTFE) sleeve with an outer diameter of ∼760 μm was mounted to one end of the wire to facilitate sample loading into the soft tube via a pipette tip and syringe, as shown in Fig. 1(a).

The wire assembly was placed inside a 15 ml Falcon tube filled with PDMS, with both ends connected to a voltage source. Applying 20 V for 15 s cured the PDMS around the wire, forming a thin, hollow PDMS soft tube. The wire with the cured PDMS tube was removed from the Falcon tube, rinsed with isopropanol, and placed in an oven at 80 °C for 15 min. To prevent damage to the PDMS tube when connecting a syringe tip for C. elegans loading, the part of the PDMS tube with the PTFE sleeve was reinforced by coating with additional PDMS solution and cured in the oven for 1 h, resulting in thicker walls around the PTFE sleeve section. The nichrome wire, immersed in the PDMS solution with the PTFE sleeve at the bottom of the Falcon tube, was then sonicated in an acetone solution for 5 min. This allowed for the gentle removal of the wire along with the PTFE sleeve from the PDMS microtube.

A modified electrothermal curing method was employed for zebrafish larvae (∼500 μm in diameter). A stainless-steel rod (120 mm length; 0.84 mm diameter) was cleaned with isopropyl alcohol. An eight-round nichrome wire coil (0.25 in. internal diameter) served as the heating element, with a voltage of 4.0 V and a current of 2.3 A applied to generate sufficient heat for curing. Once PDMS (15:1 base to curing agent ratio) was prepared, it was allowed to cure at room temperature for two hours. Then, the rod was dipped into PDMS at 1 mm/s using a motorized linear stage (Thorlabs, NRT100), heated, and cured while passing through the coil, as shown in Fig. 1(b). Upon complete emergence, the PDMS tube was carefully stripped off from the stainless-steel rod through a 0.6 mm or 22 AWG strip of a wire stripper.

The mechanical properties of the PDMS tubes were characterized by stress–strain analysis using a force tester (Mecmesin, MultiTest 0.5-dV). Young’s modulus was determined for C. elegans and zebrafish tubes by measuring the load during elongation. The data were used to ensure that the tubes could withstand the required manipulations without damage.

We used wild-type (N2 strain) worms for bright-field imaging in the adult stage. For fluorescence imaging, one transgenic model strain SJ4103 (zcls14[myo-3::GFP(mit)]) in the adult stage with GFP expressed in the mitochondria of body wall muscle cells was used in this study. It was purchased from the Caenorhabditis Genetics Center (University of Minnesota, Minnesota, MN). Worms were cultured on a nematode growth medium (NGM) with a lawn of Escherichia coli (OP50).¹⁵ Sodium hypochlorite treatment was carried out on gravid adult animals to get embryos, and then, eggs were allowed to hatch at room temperature. L1 worms were cultured at 17.5 °C for ∼70 h until the young adult stage.

Adult zebrafish (Danio rerio) were maintained in a circulating aquaculture system on a 14:10 h light:dark cycle at 28 °C. Embryos were generated by the natural spawning of WT-ABNYU in group mating and raised in an embryo culture medium as previously described.¹⁶ All protocols were approved by the New York University Abu Dhabi Institutional Animal Care and Use Committee (IACUC’s protocol ID number for zebrafish: 22–0003A3).

To capture high-resolution images of C. elegans and zebrafish larvae, we used a Nikon SMZ18 stereomicroscope with variable magnification settings (0.63–15.75×) and adjustable LED illumination for optimal contrast and clarity. The imaging system was equipped with a Nikon DS-Qi2 camera (16.25 MP), connected to a computer via USB 3.0 for real-time image capture and analysis. The NIS-Elements Advanced Research software (Nikon) was used for image acquisition and analysis. The intensity and distribution of the fluorescence signal were also analyzed using the NIS-Elements software to confirm injection results.

Scanning electron microscopy (SEM) images of the tubes were captured using a ThermoFischer Quanta3D microscope, with annotations performed using the accompanying software.

A pneumatic microinjector (Narishige, IM-300) was calibrated to deliver 4 nl of fluorescein sodium salt (Sigma, F6377-100G) at 250 μg/ml in nuclease-free H₂O. The solution was injected through a 19 μm needle made from a borosilicate glass capillary (World Precision Instruments, TW100-4). This needle was pulled to the desired tip size using a micropipette puller (Warner Instruments, PMP-102). For precision positioning during injection, it was mounted on a micromanipulator (World Precision Instruments, M3301). A Dino-Lite digital microscope (5MP Edge AM7915MZT) and the DinoCapture 2.0 software were used for side imaging during zebrafish injection.

The PDMS microtubes for C. elegans and zebrafish larvae were fabricated using the electrothermal curing methods described in the Methods and Materials section. The resulting PDMS microtube for C. elegans had an inner diameter of 78 μm and an average wall thickness of 276 μm with a wider opening (760 μm) on one end, enabling facile loading of the sample via a syringe. For zebrafish larvae, the resulting soft tube had an inner diameter of 840 μm and a wall thickness of 150–230 μm. The SEM images of both tubes are shown in Fig. S1 of the supplementary material. As can be seen in the SEM images, the wall thickness varied along the length direction since neither the wire nor the rod could be positioned vertically in the setup shown in Fig. 1. However, this wall thickness variability did not affect the tube’s functionality as our experiments showed. The tubes for C. elegans cured for 15, 25, and 35 s at 20 V showed an increased thickness with longer curing times (see the supplementary material, Fig. S2), confirming the tunability of wall thickness during the electrothermal fabrication process. To decrease the wall thickness as much as possible to enable facile stretching with subsequent imaging, a curing time of 15 s was selected for imaging C. elegans.

The Young modulus of tubes for C. elegans was 2.19 MPa, while it was 2.21 MPa for zebrafish (see the supplementary material, Figs. S3 and S4), corresponding to the standard value of PDMS. The tubes possessed the elasticity of PDMS to handle delicate specimens while maintaining structural integrity during stretching for manipulation and imaging.

The functionality of the C. elegans microtube was tested using a custom setup (Fig. 2) with two manual linear stages (Thorlabs, LT1) and fiber rotators (Thorlabs, HFR007).

An adult C. elegans was loaded into the microtube using a syringe, immobilized by stretching the tube, and rotated for whole-body imaging [Fig. 3(a)]. Images taken at various angles revealed different anatomical features, such as pharynx and muscle quadrants, demonstrating effective control of body orientation [Figs. 3(b) and 3(c)]. Importantly, the manipulation process in the soft tube did not cause significant stress or damage to the organisms, as evidenced by their normal behavior upon release.

The same setup was used for imaging zebrafish larvae, as shown in Fig. 2. A 5-dpf zebrafish was loaded into the soft tube using a pipette. After stretching, it was immobilized without anesthetization and rotated by θ = 90° for imaging (Fig. 4).

For demonstration, the zebrafish larva’s hindbrain was targeted at a 40-degree angle using a micromanipulator [Figs. 5(a) and 5(b)]. After the injection, the larvae resumed normal swimming behavior, indicating minimal stress. Fluorescein expression was visualized using a Nikon SMZ18 fluorescence stereomicroscope [Figs. 5(c) and 5(d)]. To verify injection accuracy, the location of the fluorescein signal was checked against anatomical landmarks. The tube and needle were reusable for subsequent injections.

Existing techniques, such as those using PDMS membranes³ or narrow constriction channels,⁴ often require complex setups and may necessitate organism paralysis, limiting their applicability for live studies. Our method achieves reliable immobilization and orientation control without needing anesthetics or sophisticated equipment.

The successful application to C. elegans and zebrafish larvae demonstrates the method’s versatility and potential as a valuable tool in biological and biomedical research. Future enhancements include the microinjection of C. elegans by creating extremely fine needles and thinner PDMS tube walls that would not break the needle tip. In addition, expanding this technique to imaging and microinjection of small multicellular organisms could broaden its applicability and impact.

In summary, the development of the soft PDMS microtube represents a significant advancement in manipulating and imaging of small multicellular organisms. Compared to existing micromanipulation methods, primarily in microfluidic devices, this technique offers a simple, efficient, and non-damaging method to immobilize and control the orientation of these organisms, facilitating detailed imaging in biological studies. Besides imaging, the proposed method also allowed an accurate flexible positioning for microinjection. The successful application to C. elegans and zebrafish larvae underscores its potential as a versatile biological and biomedical research tool. Future enhancements and applications of this method hold promise for further improving the study and manipulation of small-scale multicellular model organisms.

The supplementary material provides cross sections and stress–strain curves of the fabricated PDMS tubes, thickness variation of microtubes as a function of heating time, and SEM images of the glass needle.

We would like to thank Sarah Sahloul from the Micro- and Nanoscale Bioengineering Lab at NYUAD for her assistance with imaging and labeling the body parts of C. elegans. We also thank Shashi Ranjan from the Sadler Edepli Lab at NYUAD for providing zebrafish larvae and preparing glass needles. Support for Shashi Ranjan was provided by the NYUAD Faculty Research Funds AD092 and AD188 to Kirsten Sadler Edepli and NYUAD/Tamkeen fund for the zebrafish facility operation.

The authors have no conflicts to disclose.

Maksat Khobdabayev: Investigation (equal); Writing – original draft (equal). Thea Hayek: Formal analysis (equal); Investigation (equal). Salama Alnajjar: Formal analysis (equal); Investigation (equal). Layan Alkasaji: Formal analysis (equal); Investigation (equal); Methodology (equal). Ajymurat Orozaliev: Methodology (equal). Yong-Ak Song: Conceptualization (equal).

The data that support the findings of this study are available within the article and its supplementary material.