An Intraperitoneal Injection Technique in Adult Zebrafish that Minimizes Body Damage and Associated Mortality

Abstract

The adult zebrafish (Danio rerio), which is genetically accessible, is being employed as a valuable vertebrate model to study human disorders such as cardiomyopathy. Intraperitoneal (IP) injection is an important method that delivers compounds to the body for either testing therapeutic effects or generating disease models such as doxorubicin-induced cardiomyopathy (DIC). Currently, there are two methods of IP injection. Both methods have limitations when handling toxic compounds such as doxorubicin, which result in side effects manifesting as severe damage to the body shape and fish death. While these shortcomings could be overcome by extensive investigator training, a new IP injection method that has minimal side effects is desirable. Here, a unique IP injection method that is able to handle toxic compounds is reported. Consistently reduced cardiac function can result without incurring significant fish death. The technique can be easily mastered by researchers who have minimal experience with adult zebrafish.

Introduction

Zebrafish (Danio rerio) has gained attention as an experimental model for studying human diseases because this animal encompasses high gene and organ homology to humans, external fertilization, ease of genetic manipulation, and body transparency into early maturity, which facilitates a myriad of imaging applications¹. Unlike the straightforward process of delivering drugs directly to the water for zebrafish embryos and larvae, administering drugs to adult zebrafish presents a more intricate and challenging endeavor².

In adult fish, compounds can be delivered through passive drug delivery techniques, such as direct administration into the water, or through oral drug delivery methods like gavaging². Other approaches include coating fish food with the compounds and subsequently feeding the fish³, and direct administration of water-insoluble medications at a predetermined concentration, including retro-orbital or intraperitoneal injections^4,5. Intraperitoneal administration is preferred for in vivo studies of disease models due to its distinct pharmacokinetic advantages⁶. This method provides a high drug concentration and an extended half-life within the peritoneal cavity, offering an effective route for drug delivery^7,8. The approach is commonly utilized in research settings to ensure optimal drug absorption and distribution ⁹. While injection-based methods prove efficient for single delivery, prolonged and repeated injections often lead to body damage and chronic infection².

Currently, there are two methods of IP injection in adult zerbafish^4,10. However, both methods have limitations when delivering toxic compounds like doxorubicin, leading to severe damage to the body shape and fish mortality. The side effects can significantly complicate data interpretation. Although these challenges may be addressed with extensive training¹⁰, there is a clear need for a new IP injection method that minimizes side effects.

Here, our goal is to develop a new method of IP injection optimized for the effective delivery of doxorubicin into adult zebrafish, facilitating the generation of reliable doxorubicin-induced cardiomyopathy (DIC) models with minimized body damage and associated mortality.

Protocol

All procedures conducted were approved by the Mayo Clinic Institutional Animal Care and Use Committee, adhering to the standards outlined in the 'Guide for the Care and Use of Laboratory Animals' (National Academies Press, 2011). All zebrafish in the study belong to the Wild Indian Karyotype (WIK) strain. The details of the reagents and the equipment used for the study are listed in the Table of Materials.

1. Preparation and storage of doxorubicin stock solution

1. Obtain the doxorubicin stock from a commercial source.
   NOTE: Doxorubicin is light-sensitive, so acquire it in powder form and store it in opaque containers to protect it from exposure to light. Perform all steps for the Dox powder preparation within a chemical hood.
2. Completely dissolve the Dox powder in distilled water and prepare a stock solution with a final concentration of 5 mg/mL.
3. Aliquot the stock by dividing it into 1.5 mL tubes.
4. Wrap the tubes in aluminum foil to shield them from exposure to light.
5. Store the aliquot Dox solution at 4 °C for short-term storage (<1 month), or -20 °C for long-term storage¹⁰.

2. Grouping the fish according to their body weight

1. Group the fish with a BW difference of less than 10% together for subsequent injection.
   NOTE: To save effort at this phase, fish with less than a 10% BW variance are categorized as the same size.
2. Allow the fish to fast for 24 h prior to injection.
3. Anesthetize the fish using embryo water containing 0.16 mg/mL tricaine for 1 min.
4. Take the fish out of the water with tricaine and dab both sides of the fish body with clean filter paper to remove excess water.
5. Measure and record the BW of each fish, then promptly return the fish to a recovery tank filled with fresh system water.
   NOTE: Dox injection was performed on fish after reaching 3 months of age. In this study, researchers utilized 3-month to 10-month-old fishes. The BWs of mature WIK strain zebrafish may vary from 0.2 g to 0.5 g. Prolonged anesthesia lasting for more than 5 min, followed by a Dox injection, led to high fish mortality.

3. Preparation of the needle and station for injection

1. Determine the injection volume of Dox stock solution (e.g., 5 mg/mL) needed for each fish based on the average body weight to achieve the targeted dose of 20 µg/g.
2. Use the following formula to calculate the injection volume:
   
3. Add 1x Hank's Balanced Salt Solution (HBSS) to dilute the Dox solution calculated in step 1 for injection, reaching a total volume of 5 µL.
   NOTE: Utilize bulk solution for each group of fish based on their BW and include an additional 3 fish in each group to ensure that there is no shortage of solution for injection during the course of the experiment.
4. Gently tap the tube and then briefly microcentrifuge at top speed to collect the solution at room temperature for 10 s.
5. Place the prepared solution on ice and shield it from exposure to light.
6. Place a clean 100 mm Petri dish with a sponge underneath a dissecting microscope, then adjust the focus.
   NOTE: The sponge contains a 4 cm long groove. The elastic retraction of the sponge will provide the fish with support and keep the fish body in position. The sponge can be reused.
7. Equip a 10 µL micro-syringe with a 34 G beveled needle.
8. Flush the needle with 1x HBSS buffer to eliminate any bubbles and clear potential blockages from the syringe.
9. Measure 5 µL of the solution prepared in step 4 for the injection.

4. IP Dox injection procedure

1. Put the adult fish in water with 0.16 mg/mL tricaine for 1 min to induce a state of unconsciousness.
2. Position the fish in the groove of the embedded sponge with the abdomen facing upward (Figure 1A).
3. Insert the needle with an angle near 0°, starting from the midpoint of the pectoral fin towards the posterior side of the cardiac cavity (Figure 1B).
   NOTE: Avoid any contact with the heart during the procedure.
4. Direct the needle towards the tail and go beneath the silver skin.
   NOTE: Position the needle in close proximity to the silver skin, taking care to avoid any scratching or piercing.
5. Monitor the needle tip within the abdominal cavity throughout the entire operation (Figure 1C).
   NOTE: Avoid damaging the liver, intestines, swimming bladder, and other organs. Ensure that the needle reaches the end of the intestine, near the cloacal foramen.
6. Gradually and evenly dispense the 5 µL Dox solution, then slowly withdraw the needle along the original path to prevent any leakage (Figure 1D).
7. Monitor the abdominal cavity for the presence of Dox by observing a red coloration of Dox solution (Figure 1E).
8. Swiftly move the injected fish to a clean crossing tank filled with fresh system water to help the fish recuperate.
   NOTE: Between injections, flush the needle once with 1x HBSS buffer.

5. Post-injection fish management

1. Return the fish back to the system with circulation after the injection.
2. Fast all the injected fish for an additional 24 h to facilitate their recovery.
3. During the first week, keep a close eye on the fish. Remove the deceased fish as soon as possible to avoid infecting the other fish.
   NOTE: Fish fatalities within the initial 24 h are likely due to physical injuries from the injection or prolonged anesthesia. Record fish numbers to generate a survival curve.
4. Perform echocardiography to phenotype the Dox-injected fish at 56 days post-injection¹¹.
   NOTE: Ensure uniformity in conditions and procedures for the corresponding control group injected with HBSS solution.

Representative Results

Previously, two intraperitoneal (IP) methods have been employed for the administration of doxorubicin in adult zebrafish^4,10. In method I, also known as the Classic IP injection method as described by Kinkel et al.⁴, the needle was inserted at a 45° angle to the midline between the pelvic fins with the abdomen facing upward. In method II, or the Alternative IP injection method as described by Ma et al.¹⁰, the needle was inserted through the dorsal side of the fish (Figure 2A(i,ii)). By contrast, our method III has two major changes. First, the position of the injection is altered, with the abdomen facing upward and the needle directed from the midpoint of the pectoral fin toward the abdomen. The penetration site is guided by a natural hole in this area. Second, the angle of the needle is reduced, which should be close to 0° (Figure 2A(iii), Figure 1).

Comparison among three intraperitoneal injection methods
To compare the three IP injection methods, data obtained from Researcher I is presented, who did not have previous experience with zebrafish. When methods I and II were used for IP injection, significant fish mortality in the initial two weeks following IP injection was noted. In method I, death was observed three days post-injection, while in method II, it occurred after two days post-injection. 70% of the injected fish exhibited visible body damage, especially around the injection site, manifesting as drastically reduced body size and curved body shape one to two months post IP injection (Figure 2A (iv,v)). It was reasoned that the mortality and observed damage could be attributed to the toxicity of doxorubicin¹², which might leak from the opening end of the injection needle during the injection process, damaging the internal organ along the route of the injection. In method I, the tip of the injection needle could touch the gut (Figure 2A(i)), while in method II, it could touch the swim bladder (Figure 2A(ii)). It was postulated that one could prevent the tip of the needle from touching any internal organ by using method III (Figure 2A(iii)). Indeed, a significantly improved survival rate was noted, with 80% of the fish surviving at 1 month post-injection (Figure 2B). This signifies a remarkable improvement compared to the survival rates for methods I and II, which were 53.33% and 33.33%, respectively. Importantly, 80% of the survived fish exhibited a normal body shape, in contrast to those generated by methods I and II (Figure 2A(vi)). To validate the successful generation of the DIC model, cardiac function was evaluated by measuring ejection fraction percentage (EF%). Indeed, a significant reduction in EF% was observed at 56 days post-injection in the group using method III (Figure 2C).

Reproducibility of method III by different researchers
To prove that method III can be easily mastered by different researchers, 3 researchers were recruited to the study who had never worked with zebrafish before. Upon their first several tries with method III, all researchers were able to achieve 85.71%, 95%, and 83.33% survival rates 56 days after injection and a markedly decreased EF (Figure 3A,B). Of note, three researchers repeated their experiment 3 times, 3 times, and 2 times, respectively, and consistently obtained successful results. These data confirmed the repeatability of method III, allowing researchers without prior experience working with fish to generate reliable DIC models.

In one of the initial attempts using method III, researcher II tested a deviation of method III: instead of 0°, a 45° angle was used while penetrating the needle through the skin (Figure 2A(iii), dashed arrow). Only 15% of the fish survived two months after injection (Figure 3A), with 80% of the survived fish displaying considerable body damage. The data indicated that the angle of the penetration is a critical step for the success of method III.

Method III enables the establishment of a chronic DIC model
It was postulated that significantly reduced body damage might enable multiple IP injections in the same fish, mimicking the chronic DIC model in mice^13,14. Hence, method III was used to do a series of injections for 4 consecutive weeks, with 5 µg/g Dox per week (Figure 4A). In this experiment, 100% of injected fish survived, and 80% of them exhibited no signs of body damage. Importantly, a significantly reduced ejection fraction was noted at 56 dpi, indicating the successful generation of a chronic DIC model in adult zebrafish (Figure 4B).

Figure 1: Location and angle of needle insertion for the new IP injection method. (A) The location of the needle insertion. The fish is upside down, exposing the abdominal side. The head is to the right. A natural hole is located between pectoral fins that makes penetration of the skin simpler. (B) The angle of needle insertion. It is near to zero degrees to the surface of the fish. (C) The track of the needle under the skin. The tip of the needle is continuously monitored once inside the abdominal cavity. The entry point appears different from Figure 1A because of the skin distortion during the penetration process. (D) A fish after Dox is released. The abdominal cavity becomes red after releasing the Dox. (E) A fish after IP injection. There is no sign of leakage after withdrawing the needle. Please click here to view a larger version of this figure.

Figure 2: Comparison of three IP injection methods in adult zebrafish. (A) Schematics of three IP injection methods. (i-iii) Shown are schematics of the classic IP injection (method I), alternative IP injection (method II), and the new method (method III). Arrows indicate the sites of penetration and angle of the needle for three injection methods. Dashed arrow, an unsuccessful angle of needle penetration. Scale bar: 5 mm. (iv-vi) Reduced body size and curved body shape in DIC models. Shown are representative fish from DIC fish at 56 dpi. (B) Comparison of survival curves among three IP injection methods. The number of survived fish was recorded weekly. (The experiment began with a total of n = 15). (C) Cardiac function assessment of fish at 56 dpi following Dox stress. n = 6 fish in the 1x HBSS control group, and the experiment began with a total of n = 15 fish in each 3 batches injected with Dox. Values are shown as the mean ± standard error. Please click here to view a larger version of this figure.

Figure 3: Method III enables the zebrafish DIC model to be established consistently in the hands of 3 different investigators. (A) Survival curves of DIC fish using Method III by three different researchers. A high survival rate was noted in all three investigators. The numbers of live fish are recorded weekly. A low survival rate was noted for researcher II when an unsuccessful method III was used (45° needle penetration angle, as shown by a dashed arrow in Figure 2A). (B) Cardiac function assessment of DIC fish using Method III by three different researchers. In total, Researcher II, for 45° needle penetration angle, used 3 batches (9, 12, 14) fish for Dox injection. For 0° needle penetration angle n = 6 fish were employed in the 1x HBSS as shared control group. The experiment began with a total of n = 85 fish with RII (9, 12, 14), RIII (8, 6, 6), and RIV (12, 18) employed in each of the 8 batches injected with Dox by three researchers (II, III, IV), respectively. Values are shown as the mean ± standard error. Please click here to view a larger version of this figure.

Figure 4: Method III enables multiple injections of Dox in an adult zebrafish, recapitulating chronic DIC models in rodents. (A) Schematics of multiple injection model. 5 µg/g Dox (5 µL) was injected in four consecutive weeks. (B) The chronic DIC model manifests reduced EF% at 56 dpi. Values are shown as the mean ± standard error. Please click here to view a larger version of this figure.

Discussion

Different from the two existing IP injection methods^4,10, the new IP injection method is characterized by the following distinct features.Firstly, a unique needle penetration angle is used (close to zero); secondly, the needle penetrates the fish via a unique location, i.e., a natural hole on the ventral surface of a fish, which would facilitate the injection; and finally, the movement of the needle is from anterior to posterior. These adjustments effectively reduce organ injury, which results from minimized leakage of Dox during the penetration process. The needle's path is just beneath the silver skin, which can be continuously monitored by the investigator. Consequently, direct contact between the tip of the needle and the internal organs can be effectively avoided. After injection, all injected Dox stays in the abdominal cavity, and the only exit (puncture point) could be closed when the fish is awake and starts to swim forward.

Compared with the two existing IP injection methods^4,10, the new IP injection method consistently results in minimized body damage and enhanced survival rates.This statement was further underscored by the feasibility of multiple injections in the same fish for generating a chronic zebrafish DIC model. Importantly, compared to the previous two IP injection methods^4,10, the new method can be quickly mastered by researchers who have no previous experience in handling adult zebrafish. Reliable DIC models seem successfully obtained upon their first or second try in the hands of at least 4 different investigators. The success of the injection can be easily concluded within the first-week post-injection: >80%, preferably 100%, fish should be able to survive after the injection. All fish will survive until 56 dpi, when the majority of fish (>80%) should be free from obvious body damage. This is a significant improvement, because reliable DIC model using the first two IP methods can only be stabilized in the hand of a researcher after more than 3 rounds of practice with at least 10 fish in each round. This process could be time-consuming, because each practice required about 2 months to conclude on the success of his/her injection. Reducing tissue trauma not only improves the well-being of the fish but also minimizes confounding factors that could affect the outcomes of the study.

One critical step to ensure the success of the new method is the angle of penetration. As shown in Figure 2, when the angle of the penetration changed from 0° to 45°, significant fish death was noted, presumably owing to increased risk of contact between the tip of the needle and internal organs.

The primary limitation of the method is that the method cannot be used to deliver drugs to fish younger than 3 months. Because of their small body size, young fish cannot survive after the physical damage associated with the penetration of a needle. However, the method can be used in adult fish when their BW is larger than 0.15 g, including those aged from 3 months to 4 years old.

This new method would be widely used for drug delivery in adult zebrafish, especially when dealing with potentially toxic compounds such as Dox^15,16. When it comes to non-toxic compounds, all three methods should be applicable.

Materials

Name	Company	Catalog Number	Comments
10 μL NanoFil-syringe	World Precision Instruments, Inc	NANOFIL	injection tool
34 G needle	World Precision Instruments, Inc	NI34BV-2	injcetion tool
60 mm Petri dish	fisher scientific/fisherbrand	FB0875713A	placing the sponge
Dissecting microscope	Nikon	SMZ800	Injceting the Dox
Doxorubicin hydrochloride	Sigma	D1515-10MG	drug for creating DIC model
Echocardiography	VISUAL SONICS	Vevo 3100	measuring cardiac function
Foam Sponge	Jaece Industries	L800-D	placing the fish
Hank's balanced salt solution (HBBS)	Thermo Fisher	14025076	Vehicle for Dox
Microcentrifuge	southernlabware	MyFuge/C1012	collect the Dox solution
Precision Balance Scale	Torbal	AD60	Digital scales
Tricaine	Argent	MS-222	Anesthetizing fish
Tube	Eppendorf	1.5 mL	storage
vevo LAB  software	FUJIFILM VISUAL SONICS	5.6.0	quantification of the heart