Long-term Live Imaging Device for Improved Experimental Manipulation of Zebrafish Larvae

Summary

This manuscript describes the zWEDGI (zebrafish Wounding and Entrapment Device for Growth and Imaging), which is a compartmentalized device designed to orient and restrain zebrafish larvae. The design permits tail transection and long-term collection of high-resolution fluorescent microscopy images of wound healing and regeneration.

Abstract

The zebrafish larva is an important model organism for both developmental biology and wound healing. Further, the zebrafish larva is a valuable system for live high-resolution microscopic imaging of dynamic biological phenomena in space and time with cellular resolution. However, the traditional method of agarose encapsulation for live imaging can impede larval development and tissue regrowth. Therefore, this manuscript describes the zWEDGI (zebrafish Wounding and Entrapment Device for Growth and Imaging), which was designed and fabricated as a functionally compartmentalized device to orient larvae for high-resolution microscopy while permitting caudal fin transection within the device and subsequent unrestrained tail development and re-growth. This device allows for wounding and long-term imaging while maintaining viability. Given that the zWEDGI mold is 3D printed, the customizability of its geometries make it easily modified for diverse zebrafish imaging applications. Furthermore, the zWEDGI offers numerous benefits, such as access to the larva during experimentation for wounding or for the application of reagents, paralleled orientation of multiple larvae for streamlined imaging, and reusability of the device.

Introduction

The regenerative capacity of zebrafish larvae Danio rerio make them an ideal model organism for examining wound response as well as healing and regrowth^1,2,3,4. Access to an array of transgenic zebrafish lines and zebrafish's anatomical transparency further enhance their utility for in vivo studies of wound response events as well as longer-term regenerative processes⁴. Study of these biological processes using high-resolution time-lapse fluorescence microscopy therefore demands a live imaging zebrafish device that allows for high stability and minimal movement of the zebrafish larva while maintaining viability. It is key that the device allows for effective wounding while healing and regeneration occur unaffected by the device.

The standard live imaging stabilization method of embedding the larva in agarose during live imaging restricts growth and wound regeneration⁵ and may increase death rates since larvae begin to show sign of cardiac stress and tissue necrosis after four hours⁴. Therefore, removal of agarose from regions of interest is often necessary to allow normal development and regeneration⁶, exposing the larvae to potential damage as the agarose is cut away. Furthermore, with the agarose embedding technique, the user must orient the larvae in the short time before the agarose solidifies^5,6,7. Rapidly manipulating the larva not only requires skill of the user, it also risks damage to the larva. Although methods to stabilize the larva for live imaging have been described to circumvent these drawbacks, such as ridged agar wells³ or divets⁸, the use of silicone vacuum grease to create an imaging chamber with PVC piping or other materials⁶, and rotational tubing⁹, many of these methods are labor intensive, messy, often non-reusable and don't allow for environmental manipulation (drug treatments, wounding etc.) after the fish has been mounted.

Therefore, the zWEDGI device (Figure 1) was designed to overcome some of the drawbacks of agar mounting for long-term live imaging of zebrafish larvae while permitting manipulation of the specimen. The zWEDGI consists of three semi-open compartmentalized chambers (Figure 1A) to allow for loading, restraint, wounding and imaging of 2 to 4 days post-fertilization zebrafish larvae. The device is fabricated from Polydimethylsiloxane (PDMS) and placed onto the cover slip of a 60 mm glass bottom imaging dish. The design presented here was intended for wound healing studies, however the use of a modular design and standard fabrication technologies make the zWEDGI design modifiable and amenable to a variety of experimental procedures, especially for procedures that require minimal restraint with experimental manipulation and long-term imaging.

Protocol

Note: The base zWEDGI design was formulated for zebrafish larvae that are 2 to 4 days post-fertilization (dpf) and follow the guidelines of the University of Wisconsin-Madison Research Animals Resource Center.

1. Design and 3D Printing of Molds

1. Model the PDMS component of the device with desired geometries and attributes in a 3D modeling software⁵. Create an assembly of a blank mold and the PDMS part and generate a negative mold for the PDMS part by creating a cavity in the mold corresponding to the PDSM part. Save the mold as an .stl file for 3D printing (Figure 2, Step 1.1).
   NOTE: A stereolithography format (.stl) file of the mold design presented here (Figure 1) is available for download at https://morgridge.org/designs/.
2. Print molds using a photopolymer 3D printer (Figure 2, Step 1.2). Make multiple molds in a single printing, if possible, so more than one device can be molded with a single batch of PDMS.
   NOTE: The example shown was printed in Hi-Res mode with a 0.075 mm beam diameter and 0.05 mm layer thickness, using photopolymer resin⁵.
   1. Clean molds using a fine brush, denatured alcohol in a spray bottle, and compressed air to gently scrub and remove uncured resin. Remove any material from the microchannel regions.
   2. Post-cure the molds in a UV post cure apparatus for 60 min on each side as uncured resign is toxic to zebrafish larvae¹⁰.
3. Sand the cavity side of the mold with 200 grit sandpaper on a flat surface until all the sealing surfaces are in contact with the sandpaper (loading channel geometries and mold perimeter). Lightly sand with 400 and 600 grit sand paper, progressively, to produce smooth flush surfaces across all geometry facings (Figure 2, Step 1.3). Measure the depth of the cavity after sanding with a dial indicator to verify that it is close to the designed depth.
   1. Clean molds and cover discs (1 ¾ inch diameter x ¼ inch thick borosilicate glass or acrylic; one glass cover per mold) by placing in an ultrasonic cleaner filled with water for 30 min or by flushing under running water.
   2. Blow dry with compressed air and clean both the molds and covers with isopropyl alcohol and filtered compressed air. Use a clean bench as a place to fabricate the devices to minimize contamination from airborne debris. Leave the cleaned molds and glass covers in the clean bench or a covered petri dish until needed.

2. PDMS Fabrication of zWEDGI Device

1. Make the PDMS by pouring 184 silicone elastomer polydimethylsiloxane (PDMS) at a ratio of 5:1 (base to activator) into a plastic cup. Mix well for 2 min with a wooden craft stick, stirring the gel over onto itself, like kneading bread. For 5 molds, use 10 g of base and 2 g of activator.
   1. De-gas mixture in a vacuum desiccator for 25 - 45 min until all bubbles are gone.
   2. Fill the cavity of each of the 3D printed molds with approximately 0.75 mL of PDMS using a 10mL syringe until the mold slightly overflows with a meniscus (Figure 2, Step 2.1).
   3. De-gas the filled molds for 45 min to remove additional bubbles that may have formed when filling.
2. Apply a glass cover disc on top of the PDMS-filled mold, pressing the disc down at an angle to prevent bubbles from being trapped (Figure 2, Step 2.2). Allow excess PDMS to be expelled as the glass disc cover is applied.
3. Use small ratchet clamp to hold the cover disc tightly to the mold.
   NOTE: This creates a flat surface once the PDMS is cured and produces through holes where the 3D printed geometries are flush with the glass cover slip. Alternatively, to increase the number of devices that can be cured at one time, build a multi-clamp device (e.g. Figure 2, Step 2.3).
4. Cure the PDMS device in the clamped molds at 100 ^oC in an oven for 90 min (Figure 2, Step 2.4). Remove the molds from oven and allow to cool until they can be easily handled. If the clamping device is metal, remove the mold and cover disc assembly from the clamp to prevent the metal from continuing to cure the PDMS due to residual heat.
   NOTE: The device is easiest to remove from the mold while still slightly warm and not fully cured. However, once the mold is removed from the oven and the cover disc is removed, the device can sit for a couple of days before proceeding to demolding. If the device is demolded shortly after removing from the curing oven, place it in a covered petri dish to minimize contaminants while allowing it to fully cure.

3. Plasma-bonding zWEDGI to Glass Dish

1. Clamp the mold containing the PDMS device in a bench vice so that the mold's geometries are facing up, parallel to the working station bench.
   1. Start to remove the PDMS device by releasing the PDMS pull tab from the mold using flat-tipped tweezers. Work around the perimeter of the mold with the tweezers (like removing a cake out of a pan (Figure 2, Step 3.1)).
   2. Use filtered compressed air and tweezers to gently pull the device out of the mold by holding onto the pull tab and blowing air under the device. Work slowly, allowing the air to help separate the PDMS from the mold, taking special care with the tips of the tweezers around the thin tunnel sections.
2. Place the PDMS device upside down onto the inside of the cover of a glass bottomed dish so that the restraining tunnel wedges are touching the plastic (Figure 2, Step 3.2).
3. Place the dish cover with upside-down zWEDGI and the corresponding glass bottomed dish into a plasma cleaner with the inner glass facing upward (Figure 2, Step 3.3).
   1. Evacuate the plasma cleaning chamber until the pressure reaches 500 mTorr.
   2. Set radio frequency (RF) power to high. Expose the device and the dish to RF frequency for approximately 2 min. Slowly de-pressurize the chamber and return the device and the dish to a clean room hood.
4. Remove the PDMS device from dish cover. Flip over the PDMS zWEDGI to the correct orientation on the glass by positioning it carefully onto the center well of the glass bottom dish (Figure 2, Step 3.4)
   1. Using the back end of the tweezers, lightly press down on the PDMS device to ensure air bubbles are not trapped beneath. Apply pressure around the minute geometries of the micro channels and smooth the PDMS out to the edges (Figure 5C).
      NOTE: Complete contact between the PDMS and the glass ensures better adherence from plasma bonding. The zWEDGI device can be plasma bonded onto other glass bottomed dish formats, such as a glass bottomed 6-well plate (Figure 2C).
   2. Place a cover over the device dish before removing from the clean room hood.
      NOTE: Once the devices have been fixed to glass, the protocol can be paused until they are ready for use.

4. Channel Preparation and Loading Larvae

Note: General zebrafish husbandry was conducted per The Zebrafish Book, available online at http://zfin.org/zf_info/zfbook/zfbk.html. Adult zebrafish and embryos were maintained as described previously¹. The wild type AB strain was used. Follow the institution’s Animal Care Protocol for specifics regarding requirements for imaging live larvae.

1. Rinse the channels with a minimum of 100 µL 70% ethanol per channel, using a micropipette to rinse through the restraining tunnel. Remove ethanol and rinse 2 or 3 times with double distilled water. Allow to air dry.
2. Fill the channels with skim milk (at a concentration of 1% diluted in water) for 10 min at room temperature to minimize adherence of larvae to the glass coverslip of the dish. Then, gently submerge the device several times in double distilled water to rinse. Allow to air dry upside down.
   NOTE: Protocol can be paused here. This preparation of zWEDGI device can be done several days prior to use. Store covered at room temperature.
3. Prepare 1% low melting point (LMP) agarose by combining 100 µL 2% melted LMP agarose with 100 µL 2x Tricaine (0.4 mg/mL Tricaine - ethyl 3-aminobenzoate) in E3 buffer¹¹ pre-warmed to 38 ^oC. Maintain the 1% agarose/tricaine solution at 38 ^oC in a hot block to prevent gelling.
   NOTE: For multiphoton microscopy ¹¹, use either un-pigmented zebrafish variants (such as casper¹²) or maintain larvae in E3 containing 0.2 mM N-phenylthiourea to prevent pigment formation¹¹to minimize absorption of the near infrared wavelengths by the pigment.
   Caution: N-phenylthiourea is toxic. Follow the institution's rules for disposal.
4. Anesthetize larvae in E3 buffer containing 0.2 mg/mL tricaine (Tricaine/E3)¹¹. Wait until larvae are motionless and non-responsive to a touch stimulus.
5. Pre-wet the channels with a few microliters of Tricaine/E3.
   1. Pick up a single larva using a wide orifice pipette tip. Deposit the larvae into the loading channel (Figure 3A). Using a pipette tip or similar tool, orient the larva in the loading chamber such that the dorsal side faces the straighter edge of the loading chamber and the tail faces toward the restraining tunnel (Figure 3B).
   2. Carefully withdraw fluid from the wounding chamber, allowing the larva to flow into the restraining tunnel (Figure 3C). Remove most of the liquid while maintaining moisture around the larva (Figure 3D). This process can be assisted by tilting the dish slightly toward the wounding chamber.
6. Place 1% LMP agarose in Tricaine/E3 (~38 ^oC) over the larva's head, filling the loading chamber (Figure 3E). Allow agarose to solidify with the larva in the proper position. Add Tricaine/E3 to the wounding chamber as needed to maintain hydration. Repeat this loading process for the remaining 2 channels.
7. Using a syringe needle, carefully remove any agarose that seeped through the restraining tunnel into the wounding chamber (Figure 3F). Add additional tricaine/E3 either just over the agarose (for wounding, short term imaging, or wound treatment isolation) or to fill the culture dish (for long-term imaging). Replace the culture dish lid to prevent evaporation. Larvae can be imaged at this point (unwounded) or wounded.

5. Wounding and Imaging Larvae

1. Using a sterile scalpel blade, transect the tail fin posterior to the notochord¹¹ in the wounding chamber (Figure 3G, H). Add additional tricaine/E3 if needed and replace the culture dish lid.
   NOTE: Alternatively, depending of the developmental time window of interest, larvae can be wounded, permitted to recover in E3 and maintained in an incubator (28.5 ^oC) until the desired imaging time, at which point they can be loaded into channels as described above.
2. Install the zWEDGI device with anesthetized larvae onto an inverted microscope in a stage insert that will accommodate the 60 mm glass bottom dish. Locate the tail of the larva in the upper-most channel, rotating the dish as needed to get the tail in the desired position. Image as required for the specific experiment.
   Note: The zWEDGI is broadly applicable for high-resolution light microscopy, including widefield fluorescence and laser scanning microscopy. There are a number of parameter considerations when imaging zebrafish larvae, but specific imaging parameters are instrument, sample and experiment dependent. Here, the larval tail was imaged on a custom build multiphoton microscope^5,11 utilizing the following parameters: 40X long working distance water immersion objective, 890 nm laser excitation, 445/20 nm emission filter and 512 x 512 resolution.

6. End of Experiment

1. Remove the zWEDGI dish from microscope. Euthanize the larvae by placing the zWEDGI either on an ice water bath or at 4 ^oC for at least 20 min and assess for absence of heartbeat and circulation.
   NOTE: Because the larvae are maintained in separate compartments, the larvae can be individually recovered by gently pulling on the agar with forceps or pipette. The agar can be removed from the head region and the larva used for additional procedures, such as fixation for antibody staining or processed for RNA or protein extraction.
2. Following removal of the larvae and agar, clean the zWEDGI with ethanol and distilled water, as described in step 4.1 and set upside down to air dry. Store covered in a cool, dry location. Re-coat with skim milk (step 4.2) as needed prior to next use.
   NOTE: zWEDGIs can be re-used multiple times, until the PDMS begins to come away from the glass.

Results

The zWEDGI PDMS microfluidic device is a functionally compartmentalized device designed to accommodate four main functions (listed below) associated with live imaging of caudal fin wounding healing and regrowth in the zebrafish larvae. PDMS was chosen for zWEDGI fabrication because it is not only readily available and an industry standard for biocompatibility, but also works well in molds. Additionally, PDMS makes the device reusable and void of hard or sharp edges once the device is form...

Discussion

The purpose of the zWEDGI device is to capture 3D time lapse imaging by stabilizing and orienting the fish within the small working distance of a high-resolution microscope objective. While meeting these design specifications, it is also an improvement over traditional agar-based preparation for live imaging. There are three critical steps (below) in the fabrication of the zWEDGI, which, if not done correctly, can result in defective devices:

PDMS preparation (Figure 5A

Materials

List of materials used in this article

Name	Company	Catalog Number	Comments
Fabricate molds
Solidworks Professional Accedemic Research 3D modeling software	Dassault Systemes	SPX0117-01	Fisher Unitech
Viper Si2 SLA 3D printer	3D Systems Inc.	23200-902	3D Systems Inc.
Accura 60 photopolymer resin	3D Systems Inc.	24075-902	3D Systems Inc.
denatured alcohol	Sunnyside	5613735	Menards
UV post cure apparatus	3D Systems Inc.	23363-101-00	3D Systems Inc.
TouchNTuff nitrile gloves	Ansell	92-600	McMaster Carr
220B, 400B, 600 grit T414 blue-bak sandpaper	Norton	66261139359, 54, 52	MSC
borosilicate glass disc, 2" diameter	McMaster-Carr	MIL-G-47033	McMaster-Carr
ultrasonicator cleaner	Branson	1510R-MTH
isopropyl rubbing alcohol 70%	Hydrox	54845T43	McMaster-Carr
10oz clear plastic cup	WNA Masterpiece	557405	Amazon
6"craft stick	Perfect Stix	Craft WTD-500	Amazon
Name	Company	Catalog Number	Comments
Fabricate zWEDGI PDMS device
Sylgard 184 silicon elastomeric kit	Dow-Corning	4019862	Ellworth Adhesives
10mL syringe	Becton Dickinson	305219	Vitality Medical Inc
desiccator	Bel-Art Scienceware	F42027-0000	Amazon
4 in ratcheting bar clamp	Pittsburgh	68974	Harbor Freight
lab oven	Quincy Lab Inc.	20GC	Global Industrial
tweezer set	Aven	549825	McMaster-Carr
compressed air filtered nozzle	Innotech	TA-N2-2000FT	Cleanroom Supply
vacuum bench vise	Wilton Tool Group	63500	MSC Industrial
55mm glass bottom dish; 30mm micro-well #1.5 cover glass	Cellvis	D60-30-1.5-N	Cellvis
plasma cleaner	Harrick Plasma	PDC-001	Harrick Plasma
Name	Company	Catalog Number	Comments
Loading Larvae
Pipetteman, P200	Gilson	F123601
100% ethanol (diluted to 70% with water prior to use)	Pharmco-aaper	111000200
Transfer pipette	Fisherbrand	13-711-5A	Fisher Scientific
powdered skim milk		2902887	MP Biomedicals
double distilled water
N-phenylthiorurea	Sigma-Aldrich	P7629	Sigma-Aldrich
tricaine (ethyl 3-aminobenzoate)		C-FINQ-UE	Western Chemical
low melting point agarose	Sigma-Aldrich	A0701	Sigma-Aldrich
heat block (dry bath incubator)	Fisher Scientific	11-718-2	Fisher Scientific
E3 buffer
large orifice pipette tip, 200 uL	Fisherbrand	02-707-134	Fisher Scientific
General purpose pipette tip, 200 uL	Fisherbrand	21-197-8E	Fisher Scientific
#15 scalpel blade	Feather	2976	Amazon
25G syringe needle	BD	BD305122	Fisher Scientific
Name	Company	Catalog Number	Comments
Imaging
inverted microscope
Imaris imaging software	Bitplane