Live imaging of Aiptasia larvae, a model system for coral and anemone bleaching, using a simple microfluidic device

Coral reefs, and their associated diverse ecosystems, are of enormous ecological importance. In recent years, coral health has been severely impacted by environmental stressors brought on by human activity and climate change, threatening the extinction of several major reef ecosystems. Reef damage is mediated by a process called ‘coral bleaching’ where corals, sea anemones, and other cnidarians lose their photosynthetic algal symbionts (family Symbiodiniaceae) upon stress induction, resulting in drastically decreased host energy harvest and, ultimately, coral death. The mechanism by which this critical cnidarian-algal symbiosis is lost remains poorly understood. The larvae of the sea anemone, Exaiptasia pallida (commonly referred to as ‘Aiptasia’) are an attractive model organism to study this process, but they are large (∼100 mm in length, ∼75 mm in diameter), deformable, and highly motile, complicating long-term imaging and limiting study of this critical endosymbiotic relationship in live organisms. Here, we report ‘Traptasia’, a simple microfluidic device with multiple traps designed to isolate and image individual, live larvae of Aiptasia and their algal symbionts over extended time courses. Using a trap design parameterized via fluid flow simulations and polymer bead loading tests, we trapped Aiptasia larvae containing algal symbionts and demonstrated stable imaging for >10 hours. We visualized algae within Aiptasia larvae and observed algal expulsion under an environmental stressor. To our knowledge, this device is the first to enable time-lapsed, high-throughput live imaging of cnidarian larvae and their algal symbionts and, in further implementation, could provide important insights into the cellular mechanisms of cnidarian bleaching under different environmental stressors. The ‘Traptasia’ device is simple to use, requires minimal external equipment and no specialized training to operate, and can easily be adapted using the trap optimization data presented here to study a variety of large, motile organisms.


Supporting Information Text Extended Methods
Device Design and Mask Printing. Devices of trap apertures of [20,30,40,50] µm and chamber heights of [50, 70, 90] µm were designed and fabricated for bead loading and empiric optimization trials; 90 µm chamber height with 20 µm aperture were used for all subsequent Aiptasia experiments. All designs were generated in AutoCAD (Autodesk). Final triangle trap designs are available as Supplemental Resource 1 and all designs used in the study at all stages of iteration are available on an Open Science Framework project for this work. Designs were printed on 32,000 DPI transparency film (Fineline Plotting) for subsequent photolithography and device fabrication.
Photolithography. Devices were fabricated via standard soft lithography protocols (1). Briefly, single-layer devices were prepared via photolithography and subsequent replica molding into PDMS negative relief channels in the Stanford Microfluidic Foundry. For photolithography, Si 4" test-grade wafers (University Wafer) were cleaned with methanol and dried for 10 minutes before use. A 5-µm adhesion layer was applied using SU-8 2005 (MicroChem Corp.), flood exposed and post-exposure baked at 95 • C for 5 min. Subsequently, SU-8 photoresist (MicroChem Corp.) of the appropriate height was spin-coated, soft baked, exposed through the trap array transparency mask, post-exposure baked, developed and hard-baked according to the manufacturer's instructions. Full lithography details, including a detailed step-by-step instruction set, are provided in [Table S Device Fabrication. Post-hard-bake, wafers were silanized with aerosolized trichloro(1H, 1H, 2H, 2H-perfluorooctyl)silane, 97% (PFOTS, Sigma) for 10 minutes to aid PDMS transfer. PDMS (RTV 615, Momentive) was prepared according to manufacturer's instructions in a 1:10 ratio and 55 g was poured onto the master mold. The wafer was baked for 45 minutes at 80 • C. Devices were peeled off, cut, and hole-punched for 23G pin-to-tubing connections. Devices were subsequently corona-wand treated for 30 s and aligned to No. 1 coverslips (Fisher Scientific) before being baked at 80 • C for 12 hours to ensure strong bonding to the coverslip surface.
Device Operation Setup. Devices were connected in-line with a syringe pump and placed on a microscope slide mount before organism loading and imaging. The complete setup consists of: 5. similar to the inlet stopcock assembly, a luer-adapter one-way stopcock (Cole Parmer) with intermediary tubing to 1/16" tubing (Tygon, Fisher) outlet connection to de-bubble the device and collect waste.
Optionally, an off-chip bubble trap (Elveflow) for eliminating in-line bubbles connected directly to the chip inlet via additional 1/16" tubing (Tygon, Fisher) and PEEK HPLC tubing (510 µm O.D., 255 µm I.D., Zeuss) can be included for further bubble control at flow rates below 100 µL/min. A parts list is available in [Table S2].
To change buffers, load cells, or add reagents, the syringe pump was stopped, the stopcock closed and tubing disconnected upstream and syringes were exchanged. This process minimized bubble introduction into the chip. A schematic of the device setup is included in [Fig. S1].

Preparation of the Microfluidic Device.
To prevent bubbles common in long-term imaging studies using single-layer microfluidic devices, devices were surface-treated to render natively hydrophobic PDMS channel walls hydrophilic shortly before each experimental run. Devices were passivated with 5% BSA solution (BSA powder, Sigma) for 15 minutes at 35 µL/min flow using a syringe pump (Harvard Apparatus) to improve hydrophilicity and device wettability for studies conducted at low flow rates (35 µL/min); for later trials at high flow rates (100 µL/min, superior bubble mitigation), this step was unnecessary for bubble prevention. To de-bubble devices, the device outlet one-way stopcock (Cole Palmer) was turned off and the device was dead-end filled using the syringe pump at 100 µL/min flow under visual inspection until all bubbles were removed from the trap area (approximately 2 minutes). For Aiptasia runs, the syringe was exchanged to artificial seawater (ASW) and flushed through the device for 1 minute at 100 µL/min after de-bubbling to prepare for cell loading. µL/min flow (infuse-only) using a syringe pump (Pico Pump, Harvard Apparatus). Bead amounts per sample were drawn from normalized dilutions of 100,000 beads/mL, estimated using manufacturer specifications and confirmed via a hemocytometer for each bead size. Using a similar setup to that used for Aiptasia experiments, beads were loaded for ∼10 s, the device was subsequently de-bubbled, and loading was continued and allowed to stabilize for 2 minutes. In some instances (especially larger bead sizes), bead clogging was observed (see Supplemental Extended Notes) at the inlet region due to small hole punch sizes; such clogging was partially mitigated by opening and closing the outlet stopcock before flow stabilization. The microfluidic device was then imaged under brightfield illumination using an Amscope Stereoscope with an ZWO ASI-174MM camera at 10X magnification. Images were analyzed for bead occupancy in ImageJ using manual counting. Trapping efficiency was assessed by number of traps occupied per bead size and per device geometry as compared to total number of traps in the area (n=90 traps). Differential trap occupancy (number of beads per trap) was also assessed per each condition.
COMSOL Fluid Modeling, Extended. Fluid flow simulations to assess flow fields and model nutrient flow through traps with and without Aiptasia were modeled in COMSOL (COMSOL Multiphysics) using the 3D Laminar Flow module. Fluid flow through models with trap apertures of [20, 30, 40, 50] µm and chamber heights of [50, 70, 90] µm was assessed under assumptions of laminar flow. Inlet velocity was set at 100 µL/min and outlet pressure was set to 1 atm. All simulations are available under the Open Science Framework for this project. Aiptasia-occupied traps were simulated by placing a rigid ellipsoid of size 5.9 * 10 5 µm 3 1 µm away from the trap aperture center for trapped flow field analysis. For smaller chamber heights, the rigid ellipsoid was scaled such that volume was conserved (to simulate deformed Aiptasia). Further parameters are described in [Table S1]. Flow rates were extracted from the sparse simulation at the half-width trap of trap aperture (Q1) and half-width between traps (Q2) for the comparative flow calculations.

Device Loading of Aiptasia Larvae.
To perform Aiptasia imaging experiments, Aiptasia larvae were loaded directly from culture wells (∼100 µL) using 1/16" tubing (Tygon, Fisher) attached to a 1 mL syringe (Plastipak, BD) with a 23G luer-lock connector (McMaster-Carr). Lines were hung from the syringe pump vertically for 2 minutes to concentrate the motile larvae, then connected upstream of the inlet stopcock or, in early trials, directly to the device. A full protocol is described later in Supplemental Information. In trials with DCMU treatment, seawater containing 25 µM DCMU (Diuron, Sigma), an herbicide and environmental stressor, was introduced instead of normal seawater after larval loading.
Image Acquisition, Extended. Four-dimensional imaging data (time, fluorescence channels, and z-slices) of Aiptasia larvae in the microfluidic device traps were acquired on a Leica DMI6000B stand equipped with a Yokogawa CSU-10 spinning disk head, QuantEM camera (Photometrics), and ASI MS2000 motorized XY stage under 20X magnification (Leica 20X/0.7 NA multi-immersion, used with glycerine solution). Z-stacks were collected in transmitted light and chlorophyll autofluorescence channels (561 nm excitation, 405/488/561 nm dichroic and 637/37 nm emission, Semrock) under control of SlideBook 6 (Intelligent Imaging Innovations). Where noted, detailed time-series focusing on one trap were collected in the transmitted light channel only.
Image Analysis, Extended. Images were analyzed in Fiji (ImageJ) (2). For algal fluorescence quantification, images in the time series were z-projected from the z-stack by sum intensity on the chlorophyll auto-fluorescence channel and minimal intensity on transmitted light; data in Fig. 6b is generated from mean fluorescence values (arb. units) extracted from equal size ROIs in the animal, expulsion area and background per frame from sum projection image in the algal fluorescence channel for each frame. For simple visualization, mid-plane z-slices in both channels were displayed and false-colored by transmitted light (grayscale) and chlorophyll autofluorescence (Lookup Table: Green Fire Blue) to create time-course images and movies.

Open Data
Access. An open-source project repository for design files, imaging data for all trials (compressed as .avi image sequences), protocols, and additional resources, as mentioned in the text, is available at our Open Science Framework page, which can be found at OSF under [doi: 10.17605/osf.io/j2rsy].

Extended Notes
Bead Loading (Extended). During the bead capture trials displayed in Fig. 3 and Fig. S3, larger beads appeared to clog at the inlet, likely due to steric restrictions imposed by the 23G hole punch. These occasional clogs, combined with size poly-dispersity of the beads and lack of bead deformation, complicated efforts to accurately model trapping of large, deformable organisms. Therefore, these measurements can be used as approximate loading conditions for a variety of different organism sizes beyond those characterized in smaller cell literature, but should be supplemented with empirical testing with the chosen organism.
Seawater Retention (Extended). The initial 4 frames within Fig. 4 (time: 0-120 min) images were acquired before a change in room lighting; remaining images were acquired immediately following that first capture with no additional changes. Images displayed in this figure therefore contain a difference in background intensity due to this change in local lighting conditions.

Device Operation Protocol & Troubleshooting
Device Operation Protocol. We used the following protocol for conducting Aiptasia imaging experiments. This protocol could easily be translated to other organisms, if desired.
1. Set up the device as shown in [Fig. S1]. Load a syringe with desired nutrient solution (e.g. seawater) and turn on the syringe pump to 100 µL/min. Plug in the device inlet connections and open the inlet and outlet stopcocks to allow for flow through the trapping array. 3. Flow the nutrient solution for 1 min at 100 µL/min and prepare for cell loading.
4. When ready, close the inlet stopcock, stop the syringe pump, and load 50 µL cells or organisms at the desired density (∼1000 Aiptasia larvae) into the Tygon tubing connected to the syringe. Reconnect the connections, being mindful to avoid bubbles.
5. Wait up to 2 minutes for organisms to concentrate in the loading tubing upstream of the stopcock. Afterward, resume flow and open the inlet stopcock.
Other sample-loading loops and/or multiple syringe pumps in series can be used for exchanges as described in Step 4, if desired. This protocol and accompanying setup represents one cost-effective solution but is by no means meant to be restrictive or authoritative. Most importantly, stopcocks or similar mechanisms must be used during de-bubbling and loading to mitigate bubbles and organism loss during device preparation and buffer exchange in any protocol.
Bubble Reduction and Troubleshooting. In troubleshooting larval loading into the microfluidic device, we found bubble nucleation due to sample line exchanges and low flow rates to be dominant sources of reduced Aiptasia capture, trapping retention, or, in the case of massive bubble nucleation, premature larval death. We recommend the following guidelines for operation: • Flow rates for the duration of the experiment should be set as high as possible (typically >20 µL/min). High flow rates keep the chamber fully expanded at all times, preventing bubbles from traveling through PDMS. These high flow rates also aid in reducing back-swimming in motile organisms, from our observations with Aiptasia. Early trials (such as those in Fig. 4, 6, S5, S6, S9) were conducted at 35 µL/min, which had low bubble nucleation, but occasional bubble nucleation was still observed even after de-bubbling. Later trials (such as those in Fig. 5) used 100 µL/min flow rates and the full stopcock assembly as shown, and no bubble nucleation was observed after initial debubbling. We recommend this higher flow rate for ease-of-assay.
• An in-line bubble trap with a stopcock for switching reagents or input solutions should be used to eliminate in-line bubbles during sample or syringe exchanges. We recommend the system used here, but more sophisticated sample-loading loops or systems are compatible as well.
• Syringe pumps should be aligned vertically above the sample plane when loading large organisms to prevent organism settling in horizontal positions in the loading tubing. Fig. S1. Experimental Setup Overview for Aiptasia Trapping Experiments. The microfluidic device is positioned on the imaging platform and animals and reagents are introduced via a syringe pump above the device plane connected to an inlet stopcock assembly in-line with device tubing to the inlet, as shown. A similar device outlet stopcock assembly from the device outlet to waste is used for de-bubbling the device. An operating protocol is described in Supplemental Information and a parts list is available in [Table S2].          [Table S3]. Hard bake 65°C-165°C at 120°C/hr, 2.5 hr timer, auto-off Movie S1. Aiptasia larvae visualized in dish culture under a stereoscope (10 ms exposure, 14.76 s clip, shown at 10 fps).