[Methods in Molecular Biology] Chemotaxis Volume 571 || Neutrophil Motility In Vivo Using Zebrafish

BookID 157126_ChapID 10_Proof# 1 - 11/5/2009

Chapter 10

Neutrophil Motility In Vivo Using Zebrafish

Jonathan R. Mathias, Kevin B. Walters, and Anna Huttenlocher

Summary

Zebrafish have emerged as a powerful model organism to study neutrophil chemotaxis and inflammation in vivo. Studies of neutrophil chemotaxis in animal models have previously been hampered both by the limited number of specimens available for analysis and by the need for invasive procedures to perform intravital microscopy. Due to the transparency and cell permeability of zebrafish embryos these limita-tions are circumvented, and the zebrafish system is amenable to both live time-lapse imaging of neu-trophil chemotaxis and for screening of the effects of chemical compounds on the inflammatory response in vivo. Here, we describe methods to analyze neutrophil-directed migration toward wounds using both fixed embryos by myeloperoxidase activity assay, and live embryos by time-lapse microscopy. Further, methods are described for the evaluation of the effects of chemical compounds on neutrophil motility and the innate immune responses in zebrafish embryos.

Key words: Zebrafish, neutrophil, chemotaxis, myeloperoxidase activity assay, time-lapse microscopy

the zebrafish, Danio rerio, has become a powerful model organism to study cellular activities and cell migration in vivo (1). two significant advantages of the zebrafish system are (1) the large numbers of embryos that can be obtained, which facilitates quantitative studies, and (2) the optical transparency of these embryos, which enables microscopic observation without the need for surgery or invasive procedures. Another advantage of zebrafish embryos is the permeability of the embryos to chemical compounds that has enabled high-throughput screening of the effects of small molecules (2–4).

Zebrafish embryos have also recently emerged as a highly effective model system to study hematopoietic cells (5, 6) and the

1. Introduction

Tian Jin and Dale Hereld (eds.), Chemotaxis, Methods in Molecular Biology, vol. 571 DOI: 10.1007/978-1-60761-198-1_10, © Humana Press, a part of Springer Science + Business Media, LLC 2009

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152 Mathias, Walters, and Huttenlocher

BookID 157126_ChapID 10_Proof# 1 - 11/5/2009 BookID 157126_ChapID 10_Proof# 1 - 11/5/2009

immune response in vivo (7–12). Zebrafish develop neutrophils, which can be identified by their amoeboid shape and cytoplas-mic granules (13), and in fixed samples by the expression of the neutrophil-specific protein myeloperoxidase (mPO, also referred to as myeloid-specific peroxidase, mPX) (14, 15). neutrophil development beyond 2 days post fertilization (dpf) occurs within the caudal Hematopoietic tissue (cHt, see Fig. 1e), a recently described region of early hematopoietic development (16).

Fig. 1. Zebrafish embryo wounding and MPO activity assay. (a–d) Sequential images of a wound being induced in the dis-tal tailfin of a 3 dpf embryo. (a) Tailfin prior to wounding. (b) The tip of a 25-gauge needle (lower right corner) is pressed down onto the tailfin, against the dish. (c) The tailfin immediately following the wound. (d) The tailfin ~5 min after wound-ing – note that fin cells immediately around the wound have contracted and/or rounded up. (e–g) Embryos wounded and fixed at 2 h post-wound, then labeled by the MPO activity assay; individual neutrophils are indicated with arrowheads. (e) PTU-treated embryo at 3 dpf, whole body view – at this stage neutrophils are mainly found in the head, around the heart, and in the Caudal Hematopoietic Tissue (dashed box). (f) Higher magnification of boxed region in (e), dashed line outlines tailfin – note the accumulation of MPO-positive neutrophils around the wound (*). (g) 3 dpf embryo tailfin as in (f), only without PTU treatment; note pigment (arrows), which can usually be differentiated from MPO-positive neutrophils.

Neutrophil Motility In Vivo Using Zebrafish 153


Previous studies indicate that there are at least two inflam-matory leukocytes, neutrophils and macrophages, that are recruited in response to tissue wounding in larval-stage embryos (8, 9). the quantification and detection of neutrophils recruited to tissue wounding can be easily performed using methods to detect endogenous mPO activity using modifications of pre-viously described methods (7, 15). Here, we describe the use of a commercially available kit for the detection of myeloper-oxidase activity and quantification of neutrophil recruitment in fixed zebrafish embryos (9, 17). Zebrafish neutrophils can also detected in fixed embryos by using Sudan Black staining (13, 18) or in situ hybridization of mPO mRnA (14, 15). Further, we will describe methods to analyze neutrophil migration in live zebrafish embryos using time-lapse microscopy (7, 9). this pro-tocol can be performed using conventional light microscopy and differential interference contrast (DIc) to detect leukocytes, both neutrophils and macrophages, as they directionally migrate to the wound. Alternative methods to detect the specific migration of neutrophils in vivo can be performed using transgenic models of zebrafish that express GFP from the neutrophil-specific pro-moter, mPO (9, 11). We will also discuss methods that can be used to screen for the effects of chemicals on neutrophil migra-tion and inflammation in vivo.

1. Adult zebrafish (AB wild-type strain) can be purchased from the Zebrafish Resource center (Eugene, OR) or from a local pet store. However zebrafish purchased from a pet store are much more likely to have diseases due to improper husbandry.

2. Adult Fish Water: 90 mg/L sodium bicarbonate (Sigma or Aquatic Ecosystems), 50 mg/L Instant Ocean Salt (Aquatic Ecosystems), 10 mg/L calcium sulfate. Due to differences in local water sources, these parameters should be used as a guideline; in particular, the amount of Sodium Bicarbonate may vary and should be added at a concentration that yields a pH of around 7.5.

3. Small nets for handling adult fish. 4. mating chambers. mating chambers are generally clear plas-

tic boxes (0.5–1.0 L volume) with a sieve that separates adult fish from embryos after they have been laid to prevent adult fish from eating the embryos. chambers can be purchased from several sources. We recommend chambers from Aquatic Habitats, which come with dividers that keep individual fish

2. Materials

2.1. Zebrafish Maintenance and Mating



separate overnight. Alternatively the bottom of a clear plastic box can be covered with marbles, such that embryos can fall through to the bottom where they are inaccessible to adults.

5. Plastic tea strainer(s) to collect embryos. 6. Plastic 10*cm Petri Dishes, non-tissue culture. Dishes can

be reused several times. In between uses disinfect by treat-ment in a mild bleach solution overnight, followed by several washes in water (deionized water) and then drying. After hatching, embryos can stick to fresh plastic Petri dishes, which can possibly damage the embryos and cause an inflam-matory response thereby complicating the protocols in this chapter. For this reason we recommend maintaining embryos in reused dishes (which do not seem to stick to embryos) or disinfecting and rinsing fresh dishes as detailed earlier prior to their use for these protocols. Dishes used for chemical treat-ments or fixation (and the subsequent mPO activity assay) should not be reused for live embryos, as residual amounts of chemicals left on plates may damage or kill embryos.

7. Flexible plastic transfer pipettes, 7-ml total volume. 8. E3 medium for embryos: 5 mm nacl, 0.17 mm Kcl, 0.33

mm cacl2, 0.33 mm mgSO4, 10−5% methylene blue. We usually make a 60× stock of the salts and a 0.01% stock of methylene blue, both of which are stored at 4°c. 1× E3 is kept at room temperature and can be brought to pH ~7 by addition of 0.05n naOH or 1 m tris-Hcl, pH 7.6.

9. Dissecting microscope for embryo observation. 10. Incubator set at 28.5°c. 11. N-Phenylthiourea (PtU, Sigma; also known as phenylthi-

ocarbamide) can be added to E3 medium at 0.2 mm to pre-vent pigment formation. A 50× stock (10 mm) can be made in distilled water and stored at room temperature. PTU is toxic, so wear gloves at all times when handling stock solu-tion or plates containing media. For the benefit of others, be sure to indicate plates that contain PtU (e.g., we place a large green “+” sign on plates containing PtU).

12. Pronase (Sigma). Stock solution of 20 mg/ml can be aliq-uoted and stored at −20°c or −80°c. Working solutions of 1–4 mg/ml in E3 can be stored as 10–15 ml aliquots at −20°c; warm to ~30°c before using with embryos. Working solutions can be refrozen and reused several times, but with decreasing effectiveness.

1. tricaine (Ethyl 3-aminobenzoate, Sigma; also known as mESAB). Stock solution of 4 mg/ml is made by dissolving 400 mg tricaine and 1 g na2HPO4 into 100 mL deionized water; store at 4°c. Dilute to 0.1 mg/ml in E3 for working solution.

2. needle, 25 gauge.

2.2. Endogenous MPO Activity Assay



3. Phosphate-Buffered Saline (PBS). 137 mm nacl, 2.7 mm Kcl, 4.3 mm na2HPO4, 1.4 mm KH2PO4, pH 7.3. It is useful to prepare a 10× stock of this buffer for dilution of fixative and other reagents.

4. Paraformaldehyde (16%), individual vials (Electron micro-scopy Sciences). Dilute to 4% in 1× PBS (from 10× PBS stock) for fixation.

5. tween-20 (Fisher). 6. Leukocyte Peroxidase Kit (Sigma). components: Peroxi-

dase Indicator Reagent (store at 4°c), 10× trizmal, pH 6.3, buffer (store at room temperature).

7. tt buffer: 1× trizmal of pH 6.3, 0.01% tween-20. 8. 30% hydrogen peroxide (store at 4°c). 9. Dimethyl sulfoxide (DmSO) (Sigma) (if needed for com-

pound solubility). 10. Bovine serum albumin (BSA) (Sigma) (if needed for com-

pound solubility). 11. Plastic 3-cm or 6-cm Petri dishes, non-tissue culture.

1. E3 containing 0.1 mg/ml tricaine. 2. needle, 25 gauge. 3. 1% low-melt agarose in E3. 4. 3-cm plastic Petri dishes (for upright microscope). 5. 3-cm plastic Petri dishes with glass bottom (for inverted micro-

scope). We make these by drilling a hole about 1.8 cm in diam-eter in the bottom of the Petri dish. An acid-washed coverslip with a diameter of about 2.2 cm is glued in place over the hole using ultraviolet curing adhesive (norland Products, Inc.).

6. Pipette tip is used to position embryos in agarose. 7. microscope equipped with differential interference con-

trast (DIc) and/or fluorescence imaging capabilities. We use either a nikon Eclipse tE300 inverted microscope equipped with epifluorescent illumination or a Fluoview FV1000 FV10-ASW confocal laser scanning microscope, both equipped with 20× and 60× objectives.

1. maintain adult zebrafish according to established husbandry protocols (19). male fish are somewhat slimmer and have yellow underside, while females will have a white, extended belly when they are ready for mating.

2.3. Live Microscopy of Zebrafish Embryos

3. Methods

3.1. Zebrafish Mainte-nance and Mating



2. Set up crosses of adult fish on the afternoon prior to when you want embryos. Fill mating chambers with fish water, and using small nets place one male and one female in each mating chamber; cover with a lid (with holes for air) to prevent fish from jumping out overnight. the number of crosses to be set up is dependent on the number of embryos you want for experiments. General guidelines to keep in mind are that most of the time only 50% of crosses result in embryos, and that clutch sizes range from 50 to 200 embryos. many commercially available mating chambers also come with dividers to keep fish separate, thereby allow-ing one to control the time of mating, and small plastic shrubs, which can be added to provide a place for a fish to hide from a more aggressive fish.

3. the next morning, remove dividers (if included) and allow fish to mate. Zebrafish are induced to breed by the morning light turning on; male fish will aggressively swim alongside females, which will release embryos that are then fertilized by the nearby male. Embryos will fall to the bottom of the chamber, where they should be left for 5–10 min to ensure fertilization. If fish do not mate, we have found that embryo production can be helped by combining tanks (resulting in crosses of two males with two females) or by raising the inter-nal sieve to slightly immobilize the female fish.

4. After embryos are laid, remove adult fish to a separate cham-ber and collect embryos by pouring the water and embryos through a plastic tea strainer. tap embryos into a 10-cm Petri dish containing E3 (1/2–3/4 full, enough to sufficiently cover embryos), if necessary use a squirt bottle or transfer pipette filled with E3 to remove all embryos from strainer. Alternatively embryos can be moved from mating chamber to Petri dish using a transfer pipette – if this is done be sure to change E3 in dish after all embryos have been transferred. Remove and discard all debris (fish scales, feces, etc.) from dishes with a transfer pipette.

5. For larger clutches use a transfer pipette to divide embryos into additional dishes, such that no more than ~100 embryos are in each dish. Place dishes with embryos in incubator set at 28.5°c; keep embryos at this temperature at all times unless otherwise noted.

6. At around 6 h post fertilization (hpf) or later in the afternoon observe embryos using a dissecting microscope; remove and dis-card unfertilized embryos, which will have single, misshapen cells that have not divided or gone through development. Unfertilized embryos will eventually lyse and may promote bacterial growth.



7. At 24 hpf, remove and discard unfertilized or dead embryos, which will appear white and translucent. Replace E3 media by tipping dish and allowing embryos to collect at edge of plate, and then remove enough media such that embryos are still completely covered; after this add fresh E3. If embryos lack-ing pigment are desired, exchange media at this point with E3 containing 0.2 mm PtU. Wear gloves when handling media containing PtU.

8. Embryos will naturally hatch from chorions at 2.5–3 dpf; check periodically for discarded chorions, which should be removed. Replace media as needed, keeping the level at one-half to three-quarter full. If desired, chorions can be removed after 24 hpf by treatment with Pronase (see Note 1). Hatched embryos can be moved among plates using a flexible trans-fer pipette; transfer embryos gently to avoid injury, especially when expelling embryos from pipette.

1. Incubate embryos at 28.5°c to 2–4 dpf. At these stages embryos will lie flat on their sides and neutrophil develop-ment will have commenced. We prefer to use embryos at 3–4 dpf because the embryo is more “flat” than 2 dpf (due to reduction in the size of the yolk sac) and thereby easier to wound. At 5 dpf, embryos inflate swim bladders and do not rest on their sides, making media exchange and wound-ing more difficult. For statistically significant results we rec-ommend assaying at least 20 embryos per condition tested; higher numbers may be required to address the variability of neutrophil recruitment.

2. Prior to wounding, chemical compounds can be added to the media to pretreat embryos in order to make sure embryos are suffused with the compound; concentration and duration of pretreatment would need to be determined empirically (for notes on compound solubility and long-term treatments, see Note 2). to reduce the amount of compound used, embryos can be transferred to smaller dishes (e.g., 3 cm or 6 cm diam-eter); if this is done maintain lower numbers of embryos per dish (~5 embryos per ml media). If a compound is to be tested, a vehicle control plate should be set up in parallel using an equivalent number of embryos. Dilute compound (or vehicle) into E3 in a separate container without embryos prior to replacing embryo media as detailed in Subheading 3.1, step 7. Hereafter “media” will refer to E3 with or without compound.

3. Anesthetize embryos by placing in media containing 0.1 mg/ml tricaine for ~5 min at room temperature.

3.2. Endogenous MPO Activity Assay Follow-ing Wounding



4. Wound the distal tailfins (see Note 3) with the tip of a 25-gauge needle (see Fig. 1a–d). Attempt to make wounds of uniform size, as different-sized wounds will result in differing degrees of inflammatory response (see Note 4). make note of the time of wounding for accurate fixation time points.

5. Wash embryos 1–2 times in media lacking tricaine; then place at 28.5°c until fixation.

6. At desired time point (see Note 5), fix embryos in 4% para-formaldehyde/PBS for 2 h at room temperature. If embryos are floating, tween-20 can be added to 0.01% to keep embryos submerged. to reduce the amount of fixative (and subsequent reagents) used, embryos can be transferred to smaller dishes (e.g., 3 cm or 6 cm diameter) or 1.5-ml tubes for the remainder of the protocol.

7. Remove paraformaldehyde and dispose of properly. Wash fixed embryos 3 × 5 min in deionized water (see Note 6).

8. Prepare tt buffer; set aside sufficient tt buffer (e.g., 2 ml per 3 cm plate) for diluting substrate in step 10 of this sec-tion.

9. Wash fixed embryos 3 × 5 min in tt buffer; following the third wash place embryos at 37°c. We have found that equi-libration in tt buffer – especially the third wash at 37°c – is critical for efficient, optimal labeling.

10. While embryos are incubating at 37°c, dissolve Peroxidase Indicator Reagent in tt buffer at 1.5 mg/ml (see Note 7) and warm to 37°c. Substrate/tt solution should eventually take on a purple/brown color.

11. Immediately before reaction add hydrogen peroxide (to 0.015%) to substrate/tt solution and mix.

12. Remove tt from embryos, replace with substrate/tt/hydrogen peroxide mix, and place plates at 37°c. monitor development intermittently on dissecting microscope – this should be most obvious within the cHt (see Fig. 1e) and near wounded tailfins (see Fig. 1f–g).

13. When individual mPO-positive neutrophils can be dis-cerned, stop reaction by washing labeled embryos in PBS several times. Do not allow the reaction to overdevelop, as the precipitate that is produced may extend beyond cell bounda-ries and cause individual mPO-positive cells to overlap. this will hinder counting individual neutrophils that are close together, which is especially critical at the wound.

14. count number of mPO-positive cells at (or in the vicinity of) the wound for each sample (see Fig. 1f, g).

15. For long-term storage leave embryos in 4% paraformalde-hyde/PBS (or other fixative) at 4°c.



1. Prepare microscope for image acquisition and set room temperature (or closed system of microscope) at 28–30°c.

2. Place embryos in 3-cm Petri dish, in E3 containing 0.1 mg/ml tricaine to anesthetize fish. Add chemical compound for pretreatment as in Subheading 3.2, step 2. (Hereafter “media” will refer to E3 containing tricaine, with or without compound.) If the effect of the compound is reversible, it can be washed out simply by replacing the compound-containing media with fresh E3.

3. Heat low-melt agarose in microwave oven, using short bursts, just enough to melt agarose. We use 0.8–1% agarose made up in E3 and tricane is added to 0.1 mg/ml after heat-ing. cool to around room temperature. movies can also be made with embryos in media alone, but shifts in position are much more likely. Furthermore, certain compounds may not penetrate agarose, which may require keeping embryos in media (see Note 8). If embryos are not embedded in agarose, skip to step 5.

4. take media off embryos (not all – do not allow embryos to dry) and replace with agarose – enough to cover. Swirl dish to evenly distribute embryos and quickly orient embryos while agarose is still molten using a pipette tip (see Fig. 2a). For imaging of the fin it is important to try to get embryos as flat as possible on the bottom of the dish. If multiple movies are to be taken at once (i.e., with automated stage) you may want to orient embryos parallel, in same position.

5. Allow agarose to harden, fixing embryos in position. cover top of agarose with media. Embryos will remain perfectly fine in agarose for more than 12 h as long as the agarose is covered with media.

6. Orient dish on microscope stage, bring embryos into focus. If doing fluorescence microscopy using the zmPO:GFP transgenic line or other transgenic line, check GFP level of multiple embryos to select optimal embryo. Set up camera for acquisition of either DIc or fluorescence images, or both if your microscope has dual imaging capabilities. this step is done to set embryo position for quick placement after wounding.

7. Using dissecting microscope, wound ventral tailfin of the embryo using 25-gauge needle (see Fig. 2b, and Notes 3 and 4). If using an inverted microscope the needle can come in from above (imaging below). For standard micros-copy come in at ~45° angle relative to the plate and try not to disturb agarose. If the agarose becomes too distorted it may cause the fin to shift (as agarose repositions) during image acquisition.

3.3. Live Microscopy of Zebrafish Embryos After Wounding



Fig. 2. Time-lapse microscopy of Zebrafish Neutrophils. (a) Four zebrafish embryos embedded in 1% agarose in a 3.5-cm glass-bottomed Petri dish, prior to wounding. (b) A wound (arrow) was made in the ventral fin of an embedded embryo from (a) using a 25-gauge needle (as in Fig.1b) – note slight disruption of agarose to the right of the wound, where nee-dle was inserted. (c) A sequence from a movie (20× objective) taken after wounding of a zMPO:GFP embryo in the ventral fin; each column depicts three sequential time points, with 30 s in between each frame. Shown for each time point are DIC (left frame), corresponding green fluorescence (middle frame), and overlay (right frame). A macrophage (arrowhead) and a neutrophil (arrow) can be distinguished by morphology (left frame) and by GFP expression (middle frame) as they migrate towards the wound (*). Scale bar = 25 mm. (d) High magnification (60× objective with optical zoom) DIC, green fluorescence and overlay images of a polarized neutrophil migrating near a wound (arrow denotes direction of migration) made in the ventral fin of a zMPO:GFP embryo. Granules can be seen streaming near the leading edge (LE) of the cell and GFP expression clearly denotes the boundaries of the neutrophil. Scale bar = 10 mm.



8. Replace dish on microscope stage as in Step 6 and begin acquir-ing DIc and/or fluorescent images as soon after wounding as possible. We normally acquire using a 20× objective at 30 s per frame (see Fig. 2c), although shorter timespans can be used for greater resolution of single-cell features such as pseudopod extensions. movies generally run about 3 h, but can be continued to further observe the inflammatory response (see Note 9). If the goal is to observe neutrophil morphology during migration in high resolution (see Fig. 2d) or to observe the subcellular localization of a fluorescent pro-tein during migration, a 60× objective can be used, although the distance over which a neutrophil can be tracked will be much more limited. macrophages and neutrophils can be dis-tinguished by morphology using DIc microscopy (see Note 10) or by expression level of GFP if using the zmPO:GFP transgenic line (see Note 11).

9. capture images using software such as metamorph or ImageJ, which is freely available from nIH. minor shifts in embryo position can be corrected by realigning adjacent images using the “Align” tool in metamorph, with areas of pigmentation serving as convenient markers.

10. For statistical analysis of leukocyte motility, capture enough movies (minimum of N = 3 per treatment) such that 20–30 leukocytes can be accurately tracked for each treatment. We often use the “track point” function of metamorph to track cells.

11. cell-tracking data can be copied to mS Excel to facilitate analysis. Important migration characteristics such as average cell velocity and directionality index can be calculated and compared (20, 21). A convenient way to generate a figure for each movie is to use the graph function of mS Excel to generate tracks that can be overlaid onto images acquired from the microscope.

1. to remove embryos before 2.5 dpf, incubate in Pronase (1–4 mg/ml in E3) for ~5 min at room temperature, or until first embryos come out of chorions. Since prolonged treatment with pronase may damage embryos, immediately wash 3× in E3 (~20 ml per wash), during which remaining embryos should come out of chorions, which should be removed from media. Pronase working solution can be reused for addi-tional plates; remove media from embryos and replace with

4. Notes



working solution, each time attempt to limit the number of chorions in the working solution by removing as soon as the first embryos are observed to be out of chorions.

2. If solubility becomes a problem for a particular compound, DmSO can be added (up to 1%) to media. BSA can also be added to media up to 0.2% (or higher, as determined) but should not be used for extended incubations due to potential bacterial growth. Further additions to media for compound screening have also been described (4). For long-term treat-ments with compounds, one should consider omitting PtU treatment, as this may interact with some compounds and cause complications.

3. For wounding, the most convenient body section to work with is the distal tailfin, which provides a large, flat surface area that facilitates subsequent imaging. Alternatively wounds can be made in the ventral section of the tailfin (as in Fig. 2b) just below the caudal hematopoietic tissue (cHt, see Fig. 1e), which is slightly more difficult than wounding the distal tailfin but gives leukocytes a shorter distance to travel. For time-lapse microscopy it is often useful to wound in this area, as this potentially enables the visualization of random migration within the cHt in the same movie. If the zmPO:GFP trans-genic fish is being used, we recommend wounding below the cHt (see Fig. 2b) rather than in the distal tailfin, which contains several nonhematopoietic cells that misexpress the transgene and thereby hinder neutrophil tracking.

4. the size of the wound usually (but not always) corresponds to the degree of leukocyte recruitment, with smaller-sized wounds (as seen in Fig. 1c, d) resulting in less recruitment than a moderately sized “wedge” taken out the tailfin (as seen in Fig. 1f, g). transection of the tail has also been performed (11, 12, 15); however, we find that wounds of this size can lead to a large response that can impair tracking of individual cell motility. For quantitative assays using embryos that will be fixed, effort should be made to make consistently sized wounds in the same location of each embryo. to address the variability in the assay, we recommend assaying at least 20 embryos per condition tested. Further variations in leuko-cyte recruitment due to strain background or media sterility (7) should encourage individual researchers to empirically determine the size of wound that yields a reasonably consistent result prior to performing assays of chemical compounds or transient gene knock-down (18). Wound size is an essential factor to consider for time-lapse microscopy, since a large wound may cause an overwhelming leukocyte response that could hinder analysis of individual migrating cells for cell tracking.



5. We and others have previously assayed and discussed the kinetics of the neutrophil response to tailfin wounding (7, 9). It is important for individual researchers to experimen-tally determine the optimal postwound time to assess the neutrophil response. In general we have found that 2 h post wound (hpw) is a very good starting point to assay leuko-cyte recruitment, but earlier or later time points may be bet-ter depending on the size of wound. It therefore may be useful to perform multiple time points (e.g., 1 hpw, 2 hpw, 3 hpw, etc.) using similar numbers of embryos to get a more accurate assessment of the neutrophil response. If embryos are fixed at multiple time points we recommend concur-rently carrying out the fixation and washes (steps 6 and 7 of Subheading 3.2) for each sample, then leaving each sample in the first tt buffer wash (step 9 of Subheading 3.2) until the remaining steps can be performed simultaneously on all samples. Samples can be left in tt buffer overnight at 4°c if necessary, and overnight fixation at 4°c has also been reported (17). We do not recommend leaving fixed embryos in PBS for extended periods, as this seems to result in a reduction of mPO activity.

6. When doing media exchanges during mPO activity assay, take off media down to the level of the embryos while still keeping them submerged; embryos exposed to air may become slightly dried out, which might reduce mPO activity. Beyond ~4 dpf embryos inflate swim bladders, which may make media exchange difficult as embryos do not sink to the bottom of the dish (or tube), and may be accidentally taken up by the pipette. to deal with this we have found that fixing a plastic 10-ml pipette tip to the end of a transfer pipette aids media exchange by reducing the size of the opening. If this is done take special care to prevent embryos from being sucked into or stuck to the tip, which may damage the fixed embryos.

7. the Sigma Leukocyte Peroxidase kit contains single vials of Peroxidase Indicator Reagent that are aliquoted for single uses. We have adapted this kit for multiple uses of single vials of the Indicator Reagent, which consists of two solid compo-nents (one white, one purple) that are not uniformly mixed. We have found that 1.5 mg (per ml tt) of the mixed Indicator Reagent is sufficient for labeling as long as there appears to be adequate amounts of each solid component. Alternatively, an entire vial of Indicator Reagent can be dissolved in 50 ml tt, aliquoted without hydrogen peroxide and stored indefinitely at −80°c for future use.

8. Some drugs may not penetrate agarose well. We have had some good results by placing embryos on agarose, in particular



if a small well is made (e.g., by tearing out a small bit of agarose with a pipette tip) to fit the yolk sac and partially hold the embryo in place. note that while this method would work for upright microscopy, it would not work as well with an inverted microscope. Other groups have made up agarose to contain the drug of interest (22), potentially facilitating drug delivery through the agarose. Alternatively, after embedding the embryo agarose surrounding the head can be carefully peeled away, exposing the embryo directly to the drug containing media while leaving the body and fins encased in agarose. In any case, the dose and method of drug delivery to embryo is best determined empirically.

9. We generally observe the commencement of neutrophil migration toward the wound at 30 min postwounding but often it begins sooner. If neutrophil recruitment does not occur by 60 min postwounding we tend to switch to a dif ferent embryo with a fresh wound. the timing of neutrophil recruitment will vary between embryos depending partially on the size and severity of the wound induced (see Notes 4 and 5).

10. Several morphological characteristics can be utilized to iden-tify the two types of leukocyte that are recruited to a tailfin wound (9). neutrophils are usually the first cell type to migrate to the wound, and can be identified by a relatively compact amoeboid cell body that is distinctly granular in appearance; at high magnifications neutrophil granules can be seen throughout the cytoplasm (see Fig. 2d). neutrophils migrate very efficiently and with a polarized morphology (see Fig. 2c and d). Usually later in the wound response, a second cell type is seen to migrate toward wounds. Based on immunolabeling and functional observations (mathias, Dodd, Walters, unpublished observations) we believe that these cells represent a subset of (but not necessarily all) macrophages. During migration this macrophage cell type takes on an elongated morphology (see Fig. 2c) and migrates without obvious polarization and much less efficiently than neutrophils. this movement often occurs by the extension of multiple pseudopods followed by flowing of the cell body into a single pseudopod (mathias, unpublished observations). macrophages often have ingested dots of pigment that move along with them (often trailed) during migration.

11. We have found that the level of GFP expression in a leukocyte can be used to distinguish neutrophils and macrophages in the zmPO:GFP transgenic line. At 3 dpf or later, the brightest and most obvious cells are neutrophils while macrophages express much lower levels of green fluorescence protein (see Fig. 2c).



We acknowledge Ernie Dodd and Sa Kan Yoo for acquiring microscopic images used in the manuscript, and Benjamin Perrin for initial development of the protocol for time-lapse DIc imaging of zebrafish embryos.

Acknowledgments

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[Methods in Molecular Biology] Chemotaxis Volume 571 || Neutrophil Motility In Vivo Using Zebrafish

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