Live Imaging of Chemokine Receptors in Zebrafish Neutrophils During Wound Responses

NOTE: All zebrafish were kept according to the ARRIVE guidelines and UK Home Office regulations, UK Animals (Scientific Procedures) Act 1986.

1. Generation of transgenic reporter zebrafish larvae for imaging receptor trafficking in leukocytes

1. Generate Tol2-based construct for tissue specific expression of the fluorescently tagged receptor of interest. For neutrophils, use promoter sequences from the lysozyme C¹⁵ and myeloperoxidase gene¹⁶. The construct can be designed as a fusion with a single fluorescent protein (e.g. GFP), a tandem fluorescent timer (e.g., a fast-folding GFP and a slower maturing tagRFP^8,14,17) or a bicistronic expression of reporter GFP and a control membrane marker⁹ (see Discussion for considerations when choosing the approach).
   NOTE: This construct does not recapitulate endogenous levels of receptor expression but is useful for obtaining high level of receptor expression in the cell type of interest. Consult the literature on similar receptors^{3,5,6,8,9,10,14} to decide on the position of the fluorescent tag.
2. Set up a tank containing wild type adult males and females following standard husbandry practices¹⁸, separated by a barrier the day before egg spawning.
3. On the day of egg spawning, prepare transgenesis mixture for microinjection containing 12.5 ng/μL of Tol2 DNA construct and 17.5 ng/μL of transposase mRNA⁷. Lift barriers from fish tanks shortly after the lights come in the morning (this may vary in different fish facilities) and collect eggs within 15 min for mRNA injection.
   NOTE: Ensure the DNA solution is RNase free to avoid degradation of the transposase mRNA in the mixture. An option to circumvent this is to inject eggs with separate solutions of Tol2 construct and transposase.
4. Follow standard protocols for transgenesis and microinjection of zebrafish eggs¹⁹.
5. Inject 1 nL of the solution into the cell of one-cell stage embryos.
   NOTE: The expression results are more consistent when injecting inside the cell and discarding the injections that may not be clearly inside the cell. One-cell stage embryos are aimed for because of the variability of volume injection per cell when injecting 2-16 cell stage embryos. An option would be to separate the one-cell stage injections from batches of later injections, in case these have different efficacies.
6. Check the injected embryos later in the day and remove unfertilized or dead eggs to keep the clutch healthy.
7. At 3 days post-fertilization (dpf), screen larvae under a fluorescent microscope. The marker will be visible in neutrophils, particularly in the caudal hematopoietic tissue (CHT), which is rich in these cells.
   NOTE: The percentage of cells labeled varies with different constructs, but usually 20-60% of neutrophils are expected to express the construct. Lower percentages usually predict more screening at the adult stage. It is a good practice to also verify correct localization of the receptor at the membrane, with a higher-resolution imaging approach, in a sample of these embryos before growing the fish.
8. Grow positive larvae following standard husbandry practices²⁰. These represent the F0 generation.
9. At 3 months of age, screen F0 fish for founders. Cross individual fish with a non-transgenic wild type and screen their offspring at 3 dpf for the expression of the receptor by viewing under the dissecting scope. Depending on the transgenesis success, which varies with each construct, observe a percentage of positive offspring in a subset of the crosses.
   NOTE: For good transgenesis, about a third to a half of adults will give positive offspring. The percentage of offspring that is positive in a clutch varies with the copy number of transgenes inserted and, can be between 10-60%. It is helpful to keep track of mendelian ratios within the clutches to identify single insertion transgenics (these are more easily identified in F2 clutches by looking for a 50% ratio of positive larvae)²⁰.
10. Grow the positive offspring, which represent the F1 generation.
11. At 3 months of age, screen F1 adults in the same way to establish stable F2 transgenic line.
12. Perform experiments on F2 larvae after validating the neutrophil-specific expression of the transgene.
    NOTE: During the transgenesis, one may observe varying levels of receptor expression and it is advisable to keep different transgenic clutches to obtain the most appropriate expression level for the biological questions. Initial results may be obtained in F0 or F1 larvae.

2. Collecting zebrafish embryos for assessing leukocyte wound responses

1. After having established a stable transgenic reporter line, set up a cross between adult and female transgenic fish and collect eggs the next morning.
2. Grow embryos at 28.5 °C in E3 medium (or egg water¹⁸).
3. Optionally, at 24 hpf, incubate embryos in a centrifuge tube containing 50 mL of E3 medium supplemented with 0.003% bleach for 5 min. Subsequently rinse 3 times in E3 medium by letting the tube stand for a couple of minutes to allow the embryos to settle in the bottom and then decanting and replacing the medium.
   NOTE: This provides a level of control on the infection exposure of the larvae, which may affect the behavior of leukocytes during wounding.
4. After bleaching, keep embryos in E3 medium supplemented with 0.003% of 1-phenyl-2-thiourea to prevent melanin synthesis. Methylene blue, which is often used to prevent fungal infections, is not added here, to minimize tissue autofluorescence.
5. Allow larvae to hatch naturally and use at 3 dpf when neutrophils are abundant²¹.

3. Ventral fin wounding of larvae

1. Prepare larvae for wounding. Use larvae at 2.5-3.5 dpf, when abundant neutrophils are observed. Transfer larvae to E3 medium supplemented with 160-200 mg/L tricaine MS222.
   NOTE: Concentrated solutions of tricaine should be prepared and frozen in aliquots in advance and thawed on the day.
2. Leave the larvae in E3+tricaine medium for 15 min to ensure they are anesthetized. Check their responses by gently touching with a small paintbrush or similar tool.
3. Select larvae with transgenic receptor expression under a fluorescent dissecting scope.
4. Transfer larvae to a 120 mm Petri dish in E3 + tricaine for wounding. Using a sterile scalpel cut the ventral fin of the larva, while observing under the dissecting scope (Figure 1).
   NOTE: The idea is to perform a deep enough cut to cause substantial neutrophil recruitment but without cutting the vessels of the CHT. The cut is made perpendicular to the CHT axis such that the incision nearly reaches the vessels of the CHT. This takes some practice and should first be performed with supervision on larvae that are not generated for this purpose (e.g., excess larvae from another experiment).

4. Preparation of larvae for live imaging

1. Dissolve low melting point (LMP) agarose in E3 medium by heating to obtain a liquid 2% agarose solution.
2. Allow this solution to cool down to 60 °C.
   NOTE: Keep a flask or tube with agarose in an incubator at 60 °C to avoid the agarose setting between the mounting of different larvae.
3. Pipette 0.5 mL of the liquid LMP + E3 in a glass bottom dish for microscopy imaging.
4. Pipette a wounded, anesthetized zebrafish larva along with 0.5 mL of E3 + 2x Tricaine in the glass bottom dish.
5. Gently mix the two solutions to obtain a 1% LMP/1x Tricaine agarose/E3 solution, avoiding generation of bubbles. Orient the embryo laterally and gently push down so that the caudal part of the fish is as close as possible to the glass.
   NOTE: The tissue to be imaged must be as close as possible to the glass bottom when imaging with an inverted microscope. The setting of orientation for the larva must be quick so that the agarose does not set before the larva is positioned.
6. Let the agarose solution cool down and solidify for 5-10 min. Test whether the agarose is set by gently touching the agarose gel with a small paint brush or tip.
7. Once the agarose is solid, add 2 mL of E3 supplemented with 0.2 mg/mL of tricaine to the imaging plate.

5. Live confocal imaging

NOTE: Image embryos on a spinning disk confocal microscope or equivalent (Figure 2). A laser scanning microscope can also be used but the temporal resolution of the dynamics will be more limiting. Prepare the imaging settings before bringing the wounded larva, so the response can be imaged as quickly as possible after wounding. Neutrophils arrive to the wound within 5 min and receptors in the first arriving cells may internalize within this time frame. With practice it is possible to image as early as 15 min post-wounding.

1. Turn on the microscope: laser, camera and computer as per manufacturer instructions.
2. Use acquisition software to set up the imaging settings. Choose lasers for the appropriate fluorophores and approximate exposure times based on previous experiments (acquire GFP with 488 nm and tagRFP with 561 nm laser).
3. Transfer the plate with the mounted embryo, as soon as possible after agarose sets, onto the confocal imaging spinning disc platform.
4. Use the microscope eye piece to find the fish in the dish using the stage joystick.
5. Focus on the wound area, using the focusing knob.
   NOTE: To find the area of interest it may be easier to use a low magnification air objective (10x).
6. Select the field to image around the wound. Use a 30x/40x objective with high numerical aperture to obtain sufficient resolution.
7. Use the software buttons to adjust the exposure time so that the fluorescent marker can be seen with good contrast but without saturating the signal.
   NOTE: The exposure time must be as low as possible to minimize fluorescent exposure and maximize temporal resolution in the time-lapse. The laser power depends on the condition of the laser but needs to be adjusted to a level that permits low enough exposure time for dynamic imaging.
8. Use the software buttons to select the volume to image as a z-stack
9. Set up a time lapse every 30 s for the desired duration.
   NOTE: For sterile ventral fin wounds, the maximum recruitment is observed by 2-3 h.
10. Before launching the time-lapse, take a bright field snapshot to document the field of view. If possible, acquire bright field within the time-lapse movie.

6. Quantification of receptor internalization in zebrafish neutrophils

1. Record the time-interval of image acquisition and the pixel size of the image. Keep a record of how many minutes post-wounding the imaging started.
2. Open the image datasets using Fiji by dragging the image onto the software interface, select a representative frame of interest for each dataset using the time slider, e.g., at 1-1.5 h post-wounding, and save it.
3. Proceed with MATLAB to process the image dataset.
4. Create a new script and include functions for image reading (line 6 in script called ‘select_neutrophils.m’ for centroid definition in Supplementary File 1), opening (line 11 in Supplementary File 1) and manual selection of points on the image (line 12 in Supplementary File 1).
5. Open the frame of interest by running this script (Supplementary file 1), identify the neutrophils to analyze by visual inspection, click on them and record an estimation of their centroids, both in the ventral fin wound and in the CHT.
   NOTE: The non-mobilized neutrophils in the CHT serve as an internal reference for neutrophils whose receptor distribution remains constant. This allows normalization of contrast values of cells at the wound to an internal reference.
6. Proceed with segmentation of the neutrophils in each frame using active contours technique²² as described in steps below.
7. Create a function to include the metadata required for segmentation by active contours (i.e., number of iterations, bias of contour, estimated centroid etc.)²² (see ‘wound data.m’ in Supplementary File 2).
8. Create a script that calls the function ‘wound_data.m’ to input the necessary information for segmentation of each neutrophil (line 28 in script called ‘calc_contrast.m’ in Supplementary File 3).
9. Include in the script commands for image reading (line 32 in Supplementary File 3).
10. Add the generation of a black image (i.e. image where pixel values are zeros) with an equal size to the input image (line 44 in Supplementary File 3) and the definition of a square of 10 × 10 pixels around the centroid of each neutrophil (line 45 in Supplementary File 3).
11. Include the neutrophil segmentation using active contours (lines 48-49 in Supplementary file 3) and the removal of small false detected objects (line 52 in Supplementary File 3).
    NOTE: The initial segmentation contour is the square around the centroid, which evolves by active contour technique based on the pixel intensities, the number of iterations and the bias of contour. The result of segmentation is a binary mask where all pixels have value 0 apart from the neutrophil area whose pixels have value 1.
12. Include the multiplication of the segmented binary image with the original one to get the pixel intensities of the neutrophil only, with the rest of the image being not-a-number, so it does not contribute to calculations (lines 56-57 in Supplementary File 3).
13. Add the calculation of the gray-level co-occurrence matrix for each neutrophil (GLCM)^14,23 (line 61 in Supplementary File 3). GLCM is another representation of the image showing relative position of pixels in terms of pixel intensity.
14. Include the calculation of contrast of the neutrophil based on the GLCM (lines 62,65 in Supplementary File 3). The contrast metric measures differences in intensity between neighboring pixels. Pixels are compared with pixels certain distance apart, which can be adjusted empirically based on the size of local peaks in intensity. As an indication, for the images, with a pixel size of 0.389 µm, when the receptor showed vesicular distribution each bright dot was in the range of 5 pixels. Therefore, intensities were compared in pixels spaced 5 pixels apart.
15. Add commands to save the values separately for individual neutrophils in the ventral fin (line 68 in Supplementary File 3).
16. Include the calculation of the mean neutrophil contrast value from all CHT neutrophils the same way as for neutrophils at the wound (lines 72-119 in Supplementary file 3). For CHT neutrophils, call the function ‘cht_data.m’ (line 77 in Supplementary File 4).
17. Include the normalization of the contrast value of individual neutrophils at the wound to the mean contrast of CHT neutrophils calculated above in step 6.16 (i.e., division) (line 122 in Supplementary File 3). This gives a normalized contrast that reflects how ‘dotty’ the appearance of receptor is in individual responding cells relative to control non-responding cells (Figure 3 and Figure 4).
18. Run the script (Supplementary File 3) by clicking the run symbol in the software.
19. Repeat all steps for different conditions.
20. Use statistical software (see Table of Materials) to import the results for the different conditions by creating a column table, plot the results and perform statistical test to check significance of difference between the mean values.
    NOTE: The codes for the analysis can also be found in GitHub at https://github.com/LeukocyteMotionAndDynamics/ReceptorTraffic

7. Chemokine response assays in early embryos

NOTE: This is an optional side experiment that allows testing of receptor distribution changes in response to a candidate chemokine and is independent from the experiments described above concerning neutrophil expression of the receptor constructs. Differences in ligand-induced trafficking between receptors are difficult to establish with this technique as the ligand levels are saturating¹⁴. However, if one sees ligand-internalization of a receptor in this system, this can be an indication that the ligand is recognized by the receptor in instances where the ligand identity is unclear. This is useful, because expression of chemokine receptors in established cell lines such as HEK293T cells¹⁴ can be cumbersome.

1. Set up a cross of wild type fish (e.g., AB) and collect eggs the next morning shortly after lifting the separators (as described above).
2. Inject 100 pg of fluorescently tagged receptor mRNA (e.g., Cxcr1-FT), together with 100 pg of mRNA for a membrane marker (e.g., membrane CFP). Include in the mixture varying doses of mRNA for chemokine ligand.
   NOTE: As an indication 150 pg Cxcl8a mRNA gave prominent internalization of fluorescent Cxcr1-FT (see Figure 5).
3. Rinse embryos with E3 medium and incubate at 28°C.
4. At about 7 hpf, test expression of the mRNA on a fluorescent dissecting scope and select the embryos to image.
5. Prepare 0.8 % LMP agarose in advance and keep in glass tube in a heatblock at 60 ˚C. Use a glass pipette to manipulate the embryos. Gently dechorionate embryos using a pair of forceps in each hand.
6. Aspirate an individual dechorionated embryo with the glass pipette ensuring no bubbles are at the tip. Gently release the embryo into the tube of agarose allowing it to sink into the tube.
7. Aspirate the embryo from the agarose tube, collecting some liquid agarose along the way. Gently release embryo onto the center of a glass-bottomed imaging dish. Quickly rotate embryo so that the animal pole is facing the bottom of the dish (this side must be closest to the objective when using an inverted microscope).
   NOTE: One may need to readjust the orientation of embryos while the agarose is still setting.
8. After the agarose is set, supplement with 2 mL of E3 medium.
   NOTE: The embryos at this stage are very fragile in comparison to larvae and it takes some practice to dechorionate and mount. It is important to aspirate and release as gently as possible, to avoid embryo rupture.
9. Repeat the process aiming to load 3-5 embryos per dish.
10. Image embryos on an inverted confocal microscope (see Table of Materials). Use a 40x/1.3 NA oil objective to obtain high enough resolution. Visualize mCFP, sfGFP, and tagRFP with 405, 488, and 552 nm, respectively, on the scope. Adjust filters and settings to have high contrast while avoiding saturation and minimize leak through between the channels.
11. Repeat the mounting and imaging for different conditions.