Wounding Zebrafish Larval Epidermis by Laceration

Abstract

Wound healing is a critical process for maintaining the integrity of tissues, driven in large part by the active migration of cells to cover damaged regions. While the long-term tissue injury response over hours and days has been extensively studied, the rapid early migratory response of cells to injury in vivo is still being uncovered, especially in model systems such as zebrafish larvae which are ideal for live imaging with high spatiotemporal resolution. Observing these dynamics requires a wounding method that prompts a robust wound response and is compatible with immediate live imaging or other downstream applications. We have developed a procedure for wounding the epidermis in the tailfin of larval zebrafish, which we term “tissue laceration.” In this procedure, the tailfin is impaled with a glass needle that is then dragged through the tissue, which generates a full-thickness wound that elicits a dramatic migratory wound response within seconds from cells up to several hundred microns away from the wound. This procedure can be used to interrogate the processes by which epidermal cells far away from the wound are able to rapidly detect injury and respond to the wound.

Keywords

Zebrafish; wound healing; injury; epidermis; cell migration; re-epithelialization; cutaneous wound healing

Background

Wound healing is an active area of research in cell and regenerative biology, with a major focus on elucidating the mechanisms underlying early (tens of minutes) and long-term (hours and days) responses to damage. Zebrafish larvae have emerged as a key model system to study wound healing, due to their suitability for live in vivo imaging and their ease of manipulation (Enyedi et al., 2016; Mateus et al., 2012; Miskolci et al., 2019; Yoo et al., 2012). The early response of zebrafish larvae to wounding is particularly rapid and effective, and involves contraction of the wound site through an actomyosin purse string as well as active migration of epidermal cells to the site of injury, with both of these processes initiating within seconds after wounding (Gault et al., 2014). Compared to other model systems, zebrafish epidermis has the additional advantage that the purse string and migratory wound responses are spatially segregated into distinct cell types, with the outer layer of the epidermis closing by purse string contraction while the basal layer of the epidermis closes via cell migration (Gault et al., 2014). This suggests that zebrafish might be used to specifically investigate the contribution of active cell migration to wound healing at short timescales in a living animal.

However, existing techniques for injuring larval zebrafish differ in the extent to which they promote cell migration or purse string contraction. For example, transection of the larval tail is a commonly used and straightforward wounding technique (Briona and Dorsky, 2014; Franco et al., 2019; Yoo et al., 2012; Zeng et al., 2018), which elicits a wound response that appears to involve relatively more purse string contraction than cell migration (Kennard and Theriot, 2020; Mateus et al., 2012). Laser injury can produce both migration and purse-string contraction (Gault et al., 2014), but this approach requires specialized equipment, and the nature of the damage to the tissue is complex and appears to involve distinct dynamics and regulation compared to purely mechanical wounds (Datta et al., 2017; Miskolci et al., 2019; Vogel and Venugopalan, 2003). 

In order to explore the rapid migration of epidermal cells to injury, we developed a new wounding procedure, which we term “tissue laceration,” in which the larval tailfin is impaled and torn with a glass needle. This method produces more variable wound geometries than other methods, but results in a quantitative, reproducible wound response that includes significant migration of epidermal cells from up to several hundred microns away from the wound. This method does not require specialized equipment beyond what is already available in most labs that work with zebrafish, and is flexible enough to be adapted to different situations, such as at a dissection microscope or on a confocal in the middle of an acquisition. With practice, this method is a reliable way to obtain a strong and consistent migratory response to interrogate the contribution of cell migration to wound healing in vivo.

Materials and Reagents

• Solid borosilicate glass rods, 1mm diameter (e.g. Sutter BR-100-10)
• Needle puller (e.g. Sutter P-87)
• Low-melt agarose (e.g. Invitrogen Cat. No. 16520050)
• E3 medium (see Recipes)
• Scalpel handle and blades (#11-blade)
• 35mm Glass-bottom dishes (e.g. Cellvis D35-20-1.5N or D35C4-20-1.5N)
• Flame-polished Pasteur pipettes
• Embryo manipulation tools: curved pin (Fine Science Tools catalog no. 26007-03) and small spatula (catalog no. 26007-05) mounted in a holder  (catalog no. 26018-17)
• Tricaine (Sigma catalog no. E10521)
• Modeling clay
• Petri dishes (deep well 100 x 15 mm, e.g. Fisherbrand FB0875712)
• 20ml glass scintillation vials with caps
• 1.5 ml flip-cap tubes

Equipment

• Heat block for 1.5ml tubes at 38˚C
• Dissection scope (we use a Zeiss Stemi 508)
• Inverted microscope for higher-resolution imaging (we have tried this protocol with a Nikon Ti2 and a Leica DM6000B)
• Swing arm lamp, mounted near the inverted microscope (Dazor Lighting Technology catalog no. 6134)

Procedure

Glass needles

We found success with using solid borosilicate glass rods, which provided a balance between strength and flexibility. We have also tried using etched Tungsten needles (e.g. Roboz Cat. No. RS-6063), but we found that the tips were very fragile and easily broke on the glass coverslip during laceration, while glass needles were flexible and thus did not break as easily. 

Reproducing the exact tip shape with a different needle puller will be challenging; the parameters we optimized on our needle puller can be used as a starting point and are shared below. Needles that were too short had a tendency to break rather than bend on contact with the glass coverslip; needles that were too long and thin were too flexible and did not efficiently tear the tailfin. As with any micromanipulation procedure, trial and error are always required to optimize for your precise setup. 

1. Prepare a storage container for needles by rolling out two “worms” of modeling clay that can fit inside a Petri dish in parallel lines. The needles can be gently pressed into these worms to be held in place for storage.
2. Set up the pulling program on your needle puller. Precise needle pulling parameters are not easy to reproduce between machines due to differences in filament properties, local humidity, and many other factors. However, as a rough starting point, we pulled needles on a Sutter Instruments P-87 needle puller (Flaming-Brown type) installed with a 3x3 mm box filament and a ramp calibration value of 665 using the following parameters: Heat 665 (set to ramp value); Pull 55; Velocity 60; Time 250; pressure 500. The needle was formed in a single pull, which took at least 10 seconds of heating--longer than a pull for a typical microinjection needle.
3. Insert borosilicate rods into the needle puller and pull according to the instructions of your needle puller. Remove needles and place in the needle storage container.
4. Needles were not sterilized beyond the heat of the filament used when pulling the needle, and were reused for several weeks, or until tissue debris sticks to the needle or if the solution contained poorly soluble drugs such as blebbistatin, which might adsorb to the glass. Generally needles broke from use within this timeframe.

E3 in low-melt agarose

1. Prepare a 60x stock solution of E3 (see Recipes).
2. Dilute E3 to 1X with RO water and add low-melt agarose to a concentration of 1.2% w/v. 1.2% was chosen to tolerate some dilution during the mounting process while maintaining sufficient rigidity. 2% agarose is too concentrated and can crush larvae upon gelation. 
3. Microwave the E3-agarose mixture until fully dissolved, using low to medium power to prevent boiling over.
4. Divide into aliquots of ~6-7ml (we used 20ml glass scintillation vials with screw caps but other microwaveable containers may be used), which can be stored for several months at 4˚C
5. Prior to experiments, re-melt an aliquot in the microwave using short pulses of a few seconds interspersed with stirring until agarose is clearly dissolved. Caution, the vial can get quite hot!
6. Pipet 720µl into 1.5ml tubes, and keep these tubes at 38-42˚C to keep agarose from solidifying. These working vials can be used for up to 1 week, after which time the agarose near the top of the tube starts to polymerize.

Zebrafish

Zebrafish from 24-120 hpf have been successfully wounded with this protocol.

1. Maintain zebrafish stocks according to standard procedures (Westerfield, 2007) and guidance from your institution’s department of comparative medicine or other office responsible for welfare of research animals.
2. The afternoon before embryos are to be fertilized, place 1-2 male and 1-3 female zebrafish of breeding age (at least 3 months old depending on rearing conditions) into a spawning tank. Fish will spawn the following morning after lights come on in the zebrafish facility.
3. Return adults to their housing after spawning and collect fertilized eggs by pouring the spawning water through a mesh strainer and rinsing the eggs into a petri dish with system water.
4. Split embryos into multiple petri dishes of system water at a density of no more than 60 embryos per deep-welled petri dish or 50 per regular petri dish. Inspect embryos at the end of each day, including the day of fertilization, to remove dead embryos and prevent fungal growth. Transferring embryos into fresh system water can be beneficial for minimizing microbial growth. Antifungal agents like methylene blue can also be added, but these agents will also frequently stain cell types within the larval epidermis, which can be undesirable for downstream imaging.
5. Screen embryos for desired genetic phenotypes (e.g. fluorescent markers) according to your specifications. 

Mounting

For most of the work in this paper, we opted for a quick and flexible procedure of mounting in agarose followed by cutting a window out of the agarose to allow access to the tailfin with a microneedle. Alternatively, devices like the zWEDGI can be used for immobilization of the larva while the tail remains free (Huemer et al., 2017, 2016). These devices help with consistent mounting of the larvae in the correct position, and also allow independent control of the composition of the media on the anterior and posterior side of the larvae, but require more time devoted upfront to assembly. Detailed protocols for use of these devices have been previously published (Huemer et al., 2017).

1. If the larvae are still in their chorions, dechorionate larvae and allow to straighten for a few minutes, then transfer them with a fire-polished glass Pasteur pipette to a Petri dish containing E3 + Tricaine. Allow a few minutes to anesthetize larvae, which can be confirmed with a gentle puff from a pipette or a gentle tap with a manipulating needle.
2. While larvae are being anesthetized, add 30µl of 25X Tricaine solution to the side of one of the aliquots of E3 + Low Melt Agarose and flick gently to mix without introducing bubbles. Mark this aliquot with a felt tip pen and throw it out at the end of the day.
3. Gather anesthetized larvae in the Pasteur pipette and transfer into the cap of the marked tube of agarose. Working quickly, carefully remove any excess liquid from the fish and then close the cap and invert the tube to suspend the larvae in the agarose. Flip the tube right side up and check that all larvae are in the agarose, as opposed to stuck to the walls or still in the cap--it may require shaking the tube to get the larvae into the main solution of agarose.
4. Gather the larvae and agarose using the Pasteur pipette and pipette ~500µl of solution into the well of a 35mm glass-bottom dish. Use the curved pin manipulator tool to position and mount the fish. Position the tailfins near the center of the dish to allow maximum room for maneuvering, and mount the larva on its right side so that the tailfin is in-plane with the coverslip; this requires practice and patience. Press gently on the larva on the yolk sac and yolk extension using the side of the curved manipulator to move it around in the agar. Avoid touching the larva with the point of the curved pin, which can damage the larva. Also avoid directly touching the tip tailfin as much as possible, to prevent inadvertent damage before imaging. The large size of the yolk sac will tend to cause larvae to tilt relative to the coverslip, so larvae must frequently be prodded and re-oriented until the agar starts to set, roughly a few minutes after being placed in the dish, depending on ambient temperature. You can tell the agarose is starting to set when the surface holds deformations. We typically mount 4 larvae at a time to ensure consistent mounting, though at first only one or two should be attempted at a time, and with practice more larvae can be simultaneously mounted. Larvae that are successfully mounted in a plane parallel to the coverslip will require a smaller imaging volume and therefore lead to lower phototoxicity and easier data handling. 
5. Once the agarose starts to solidify, leave it a few more minutes to fully solidify (indicated by an opaque appearance), then cover it in E3 + Tricaine solution or other solution of interest. 
6. Using a #11 scalpel blade, cut a window around the tailfin of the larvae. This blade was chosen in order to have a clear view of the tip to avoid prematurely damaging the tailfin. 
   1. Using the tip of the blade, drag it through the agarose to make two full-thickness cuts through the agarose parallel to the A-P axis of the larvae on either side. These cuts should extend anteriorly to about halfway between the tailfin and the yolk extension, but can extend less; if the cuts extend too far up the fish, then the pumping of blood flow can cause periodic movement of the tailfin that can cause motion artifacts during imaging.
   2. Next make a cut perpendicular to the AP axis, over the trunks of the larvae, in order to join the two previous cuts. Care must be taken not to injure the larva: carefully lift the blade up and over each larva, staying as close as possible to the larva, and then back down to touch the coverslip on the other side.
   3. Finally, make a cut posterior to the tailfin and parallel to the third cut, to completely cut out a rectangle of agarose from around the tailfin.
   4. Using a scooping manipulator tool (Fine Science Tools catalog no. 26007-05), insert the scoop between two larvae in the cut above the trunk and gently pry the agarose off of the tailfin and out of the dish. If the agarose is concentrated enough, the whole block should come out evenly. Sometimes this procedure breaks the agarose into small pieces, which should be completely removed to avoid interference with the needle during wounding.
   5. Using the curved manipulator tool, trace around the tailfin to check for residual agarose: if all agarose is removed, tracing around should not cause the tailfin to move, while residual agarose will catch the tailfin and cause it to bend. Generally tracing around the tailfin will also serve to fully remove the residual agarose.

Wounding

This is also a flexible procedure, which can be done several ways: first, wounding on the scope allows for imaging the same larvae immediately before and after wounding, which is necessary for observing the events within the first minute or so after wounding. With practice this can be done quickly and reproducibly by hand. Finally, a sample can be wounded manually at a dissection scope and then moved to the confocal for imaging. This approach has the advantage of increased visibility and control provided by a dissection scope, at the expense of missing the events in the first few minutes after wounding.

Alternatively, a micromanipulator can also be used to hold the needle in place, which confers greater stability to the needle at the expense of reduced control (depending on how elaborate your micromanipulator controller is) and increased setup and alignment time. This protocol discusses the manual wounding strategies in depth, and briefly touches on some considerations for micromanipulator wounding.

Manual laceration at a dissection scope

1. Take a needle and hold like a pen; your middle finger as well as fingers from your other hand can be used for further stabilization
2. With the needle, impale the larvae at a position dorsal or ventral to the terminus of the notochord--e.g. see Fig. 1A in (Kennard and Theriot, 2020)--and drag the needle in a posterior direction, tearing the tailfin. 
   1. In subsequent investigation, we have found that pulling the needle at a ~45 degree angle in the X-Y plane, following the orientation of the actinotrichia, produces a cleaner wound that is more likely to be consistent from larva to larva, and less likely to rip a large chunk out of the tailfin.
3. Repeat this laceration at a position on the other side of the notochord, so that there are two tears in the tailfin that are ideally symmetric about the dorsal/ventral axis.
4. The larva is now ready for imaging or other downstream analysis of the wound response.
5. Note: This procedure can also be used with unmounted (anesthetized) larvae in a petri dish for higher throughput. In this case, holding the curved manipulator tool (Fine Science Tools catalog no. 26007-03) in your non-dominant hand, rest the point of the tool above (dorsal to) the larva and lower it until it is securely pinning the larva against the bottom of the dish just posterior to the yolk sac. Holding the needle in your dominant hand, wound the larva as described; the wounding procedure will tug the larva in the posterior direction, which will be resisted by the manipulator tool. With practice this can be done in just a few seconds and allow wounding many unmounted larvae in a petri dish in a short period of time. 

Manual laceration at an inverted microscope

This procedure is done on an inverted scope to allow for ease of access; it has been practiced successfully on both a Leica DM6000B and a Nikon Ti2. This procedure is of course easier with lower magnification lenses, though it has been successfully and routinely applied up to 60X magnification. It is highly recommended to practice at 10X until comfortable.

1. If desired, acquire a few images of the tailfin prior to injury, then stop or pause the acquisition for wounding. Make sure to select the eyepieces in the optical path rather than the camera for the next steps. Caution: If using a confocal, make sure your lasers are off or your laser safety interlock is functioning properly to prevent inadvertent exposure of your eyes to direct laser light through the eyepieces.
2. Move the transmitted light turret out of the optical path to provide more room for manipulation; to provide light, an external lamp on a swing arm stand can be moved in position in the optical path (resting on some part of the turret if required). Using this light source has the additional advantage of illuminating the sample like a low-NA condenser, which provides a greater depth of field for wounding than is typically used for acquisition.
3. Hold a needle in one hand between thumb and the index and middle fingers. Move your arms over the sample, positioning the needle so it is in the optical path, being careful not to block the optical path with your fingers. Bringing your other hand in from the other side of the sample, you can further stabilize the needle using your other fingers, as well as resting your hands on stable parts of the microscope body. Make sure that there is no tension in your hands, to avoid excessive shaking of your fingers.
   1. Depending on the optical setup, it may be helpful to bring the needle in from the front or the back of the sample.
   2. In general, the needle needs to be positioned at an angle to prevent your hands from blocking the optical path. However, the effectiveness of tissue impalement and injury appears to be greater when the needle is as close to vertical as is feasible.
4. Carefully bring the needle into focus. First observe the position of the needle relative to the sample with the naked eye to make sure you are holding it close to the larva, then carefully reposition your body to look through the eyepieces without moving your hands too much. Gradually lower the needle towards the sample, gently swaying the needle back and forth across the field of view to provide a visual cue of when the needle is coming into focus. Often it is helpful to center a location further up the shaft of the needle within the optical path, rather than the tip of the needle; as the needle lowers into focus, you will be unlikely to miss the whole shaft, whereas a small error in positioning of the tip might cause you to move the needle outside of the field of view, or accidentally impale the larva prematurely. On the other hand, if you focus on a point too far up the shaft, you may risk lowering the needle too far and breaking the tip.
5. Once the tip of the needle is in focus and in the field of view, proceed to lacerate the tailfin as described above (“Manual wounding at a dissection scope”). Hand tremor is more noticeable in this setup than with the dissection scope, and it takes practice to impale with the same degree of control afforded under the dissection scope. Often the majority of time is spent positioning the needle “close enough” (given the degree of hand tremor), and then committing to that position and impaling the tailfin, regardless of what happens next. In practice, we have found that many interesting events following wound healing are robust to variation in the geometry of the wound.
6. After laceration, swing the lamp out of the optical path, replace the transmitted light turret, and switch the optical path back to the camera rather than the eyepieces, and resume image acquisition. 
7. Movement of the tailfin during acquisition is a greater problem in this procedure than in typical zebrafish imaging where the entire larva is embedded in agarose. Especially at higher magnifications, the larva may have shifted relative to its position prior to wounding (in X, Y, and Z). This is especially true in the first few frames as the tailfin “relaxes” back to its original position after being tugged during the wounding procedure. Re-focusing of the sample after wounding may be necessary. If possible, it can be beneficial to set up a larger volume in Z during acquisition than necessary, to accommodate such sample movement. Additional image registration approaches can also help to correct for this movement post hoc (Kennard and Theriot, 2020).

Wounding with a micromanipulator

The procedure above can also be carried out with the needle placed in a micromanipulator. This approach will greatly reduce imprecision due to hand tremor. However, micromanipulators require practice and care to align properly in the optical path prior to imaging, and can add a lot of extra setup time to an experiment. Micromanipulators also present additional opportunities to accidentally break a needle during sample exchange, which requires realignment of the setup with a new needle. Furthermore, most micromanipulators are designed for small precise motions, rather than the larger, forceful motions needed to impale and tear the tailfin. Although micromanipulators can be positioned precisely, they nevertheless do not guarantee precise wound positioning because the tailfin is quite flexible and does not stay in place as it is tugged during laceration. We have found that the manual procedures described above are a better fit for our experimental needs.

If, despite these caveats, laceration with a micromanipulator is desired, we have found in preliminary trials that the 3-axis hydraulic joystick type manipulator controller (Narishige MO-202U) to provide enough degrees of freedom and range of motion with intuitive control for effective laceration, although an even larger range of motion would be ideal.

Notes

Reproducibility

Although with practice laceration can produce consistent results, this technique does not offer as much control over the position and geometry of the wound as other methods, such as scalpel wounding or laser wounding. In our experience we find that this can be an advantage, since if the wound location is controlled too precisely, biological differences in e.g. anterior-posterior developmental patterning can lead to systematic biases in wound response from location to location (cf (Lopez-Baez et al., 2018) figure 1 supplement 1e). We have found that the response of cells not immediately adjacent to a laceration wound (~20-300µm away) is largely consistent despite variation in wound geometry (Kennard and Theriot, 2020). Thus we believe laceration is well-suited for interrogating the behavior of these more-distant cells.

Potential Issues

One of the most common failure modes is that the tip of the needle breaks against the coverslip due to too much pressure being applied on the needle. This can lead to glass fragments remaining in the field of view during your acquisition, potentially obscuring events of interest. If not much of the tip is broken off, one may be tempted to continue to use the needle. However, we find that these blunter needles are more likely to break again because the remaining portion of the tip is thicker and more brittle than the part that broke off, and with each subsequent break the tip becomes larger and less capable of impaling the tissue and producing a clean tear. We recommend having several backup needles on hand so one can quickly dispose of the broken needle in a sharps bin and carry on with a fresh needle.

Recipes

E3 + Tricaine

1. E3 was prepared as a 60X stock according to the cited protocol, without autoclaving or the addition of methylene blue (“E3 medium,” 2008). It was kept for several weeks at room temperature.
2. A 25X Tricaine stock solution (4 mg/ml stock concentration) was prepared by dissolving Tricaine into water and adding 2% (v/v) of 1M Tris, pH 9. Additional Tris solution was titrated in to bring solution to a final pH of 7. This stock solution was dated and stored in the fridge for up to 6 months. 
3. On the day of imaging, a solution of 1X E3 + Tricaine (referred to as E3 + Tricaine) was freshly prepared by diluting the 60X E3 stock and the 25X Tricaine stock to 1X with milliQ water. This solution was discarded at the end of each day.

Acknowledgements

AK and EL were supported by an NIGMS training grant (T32GM008294). EL was also supported by an NSF GRFP (DGE-1656518). JAT acknowledges support from the Howard Hughes Medical Institute and the Washington Research Foundation.

Competing Interests

The authors declare that they have no competing interests

Ethics

Work with zebrafish was approved by the University of Washington Institutional Animal Care and Use Committee, protocol number 4427–01

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