Intramyocardial Injection for the Study of Cardiac Lymphatic Function in Zebrafish

Summary

The present protocol allows efficient and stable delivery of fluorescent microspheres (MS) and quantum dots (QD) into the myocardium of live fish that can be tracked (traced) over time.

Abstract

Zebrafish have proved to be an important model for studying cardiovascular formation and function during postembryonic development and regeneration. The present protocol describes a method for injecting fluorescent tracers into the zebrafish myocardium to study interstitial fluid and debris uptake into cardiac lymphatic vessels. To do so, microspheres (200 nm diameter) and quantum dots (<10 nm diameter) are introduced into the myocardium of live zebrafish, which can be tracked using ex vivo confocal microscopy. These tracers are then tracked intermittently over several hours to follow clearance from the myocardium into cardiac lymphatic vessels. Quantum dots are transported through cardiac lymphatic vessels away from the heart, while larger microspheres remain at the injection site for over three weeks. This method of intramyocardial injection can be extended to other uses, including the injection of encapsulated MS or hydrogels to locally release cells, proteins, or compounds of interest to a targeted region of the heart.

Introduction

The lymphatic system is essential for maintaining tissue-fluid balance, modulation of the immune response following injury, and absorption of lipids in the gut¹. Accumulating evidence supports the broad roles of the lymphatic system in various disease and developmental contexts. However, mechanistic studies are hampered because lymphatic vessels can be hard to visualize, and their functionality can be uncertain. Early imaging techniques relied on the natural ability of the lymphatic system to interstitially absorb injected tracers, and then transport them through the lymphatic vessel network, allowing detection and visualization¹. Not only can this method be used to visualize the lymphatics, but it can also be used to quantify their ability to uptake fluid and macromolecules from the tissue.

The vast lymphatic network also encompasses the cardiac lymphatic system, which has been shown to play an integral role in zebrafish regeneration^2,3,4. Understanding the differences and similarities in lymphatic function across different species is crucial to utilizing this knowledge clinically. Therefore, there is a need to explore the technologies that can measure and visualize lymphatic function across different model organisms^5,6. Lymphatics are blunt-end vessels that transport fluid in one direction, away from the tissue⁷. Intramyocardial injection of fluorescent dyes is required to observe lymphatic drainage from cardiac tissue. Intramyocardial injections have also been used clinically and in pre-clinical mammalian models to transplant stem and progenitor cells or exogenous compounds such as hydrogel to test for the improvement of heart function after myocardial infarction^8,9,10. Zebrafish intramyocardial injection has not been described in detail, which has limited the use of such experimental approaches to the zebrafish heart.

Injections into the zebrafish's pericardial space and the systemic blood flow within the lumen of the heart have been described in detail previously^11,12, and successful intramyocardial injection of fluorescent tracers in adult zebrafish has been reported². The present article provides a detailed protocol for carrying out intramyocardial injections in adult zebrafish. Several transgenic zebrafish lines can identify lymphatic vessels; however, there is a need to explore approaches to understanding lymphatic drainage or to visualize lymphatics in the absence of transgenic markers. Fluorescent tracers, microspheres (MS), and quantum dots (QD) are used here to visualize the injection site and fluid flow into the cardiac lymphatics. QD are fluorescent nanocrystals of <10 nm in diameter whose optical properties can be tuned and adapted to serve many biomedical applications^13,14. QD are readily taken up by lymphatic vessels but not by blood vasculature when injected interstitially^15,16. MS are fluorescently coated polystyrene beads of approximately 200 nm in diameter¹⁵. As such, MS are considerably larger than QD and significantly more persistent when injected into the myocardium, allowing consistent identification of the injection site. This method is useful to study lymphatic function during cardiac regeneration but can be adapted to study various aspects of cardiac biology using the stable localized introduction of coated beads, hydrogels, or cell preparations.

Protocol

All animal procedures were approved by the Institutional Animal Care and Use Committee at Weill Cornell Medicine (protocol 2020-0027) and followed proper guidelines. The following experiments were performed with male and female AB wild-type zebrafish aged 14-to-20-months post fertilization for adults, and 35-days post fertilization for juveniles.

1. Needle pulling and reagent preparation

1. Pull a 1.2 mm OD (outer diameter) standard borosilicate glass capillary using a needle puller (Figure 1A,B) (see Table of Materials), into two needles using optimal settings. In this case, heat 525; pull 65; velocity 60; time 250.
2. Vortex commercially obtained stock colloidal solutions of MS (1% solids in water) and QD (2 µM, see Table of Materials).
   NOTE: The MS colloidal solution is white in color and easily seen when injected.
3. Filter 500 µL of MS stock colloidal solution through a 0.45 µm syringe filter (see Table of Materials) to minimize clogging the needle.
4. To prepare the working colloidal solution of only MS, dilute 100 µL of the filtered MS colloidal solution with 100 µL of 1x PBS (Phosphate buffered saline).
5. To prepare the working colloidal solution of MS and QD, mix 100 µL of the filtered MS colloidal solution with 100 µL of the QD colloidal solution.
   NOTE: Depending on the experiment, MS, and QD can be mixed or injected individually.
6. Prepare and arrange the necessary reagents and equipment: stereoscope, wet sponge, micromanipulator, injector, 20 µL pipette, tricaine solution, straight iridectomy scissors, forceps, pulled needles, microloader femtotips, recovery tanks, and nets as shown in Figure 1D (see Table of Materials).
7. Turn on the injector air supply and set up the gated injection pulse rather than the timed injection pulse.
   ​NOTE: For the instrumental details, please see Table of Materials. Some instruments require the injection pedal to be inserted into the gated input port on the instrument's back side.

2. Injection station preparation and zebrafish preparation

1. Turn on the dissecting microscope and adjust the focus.
2. With microloader pipette tips, load the control solution (e.g., 0.05% phenol red in 1x PBS).
3. Insert the needle into the needle holder of the microinjector.
4. Under the dissecting microscope, cut the pulled needle at 1 mm from the tip using forceps.
   NOTE: Cutting the needle as narrow as possible is optimal for piercing the myocardium. However, if the needle is too narrow, it may bend when trying to penetrate the cardiac tissue.
5. Set the appropriate injection pressure to around 0.50 kPa/7.3 psi, and the balance pressure to around 0 psi so there is no liquid retraction into the needle (Figure 1C).
6. Once the desired pressure is reached, test the injection setup, and record the time taken to inject a bolus of solution into mineral oil that is less than 1 mm in diameter (<0.5 µL).
   NOTE: Ideally, the injection must release the fluid at approximately 50 nL/s to give an approximate bolus diameter of 0.8 mm in 5 s. Depending on the tip diameter of the needle, adjustments in time and pressure can be made to inject 0.25-0.3 µL into a single fish over a 5 s injection. Higher injection pressure, as opposed to a longer injection time, is desirable to ensure that the high interstitial tissue pressure can be overcome during the injection.
7. Load the needle with the selected volume (e.g., 10-15 µL, Figure 1B) of injection solution and repeat steps 2.5-2.6.
   NOTE: Gated injections will allow one to inject for as long as the foot pedal is pressed, allowing for adjustments of the tip position to be made until the right intratissue injection site is found.
8. Insert the needle loaded with injection solution into the needle holder of the microinjector.

3. Injection

1. Prepare a grooved sponge by carving a fish-like outline in the middle.
2. Prepare the working concentration of tricaine by diluting 4.2 mL of stock solution (4 mg/mL) into 100 mL of fish facility water (see Table of Materials) in a crystallizing dish.
3. Anesthetize the zebrafish in tricaine solution until the movement of the gills is reduced and it has stopped swimming.
   NOTE: Pinch the tail to confirm the loss of response in anesthetized zebrafish.
4. Position the zebrafish with its ventral side facing the objective lens in the moistened grooved sponge under the dissecting microscope (Figure 2A and Figure 3A).
5. Use iridectomy scissors to make a small transverse cut at the level of the pectoral fins and open the chest cavity (Supplementary Video 1).
6. With forceps, gently peel away the pericardium to expose the apex of the heart (Figure 3B).
7. With the loaded needle holder in hand, direct the needle toward the apex of the ventricle at an acute angle of <30° to the body axis.
8. Insert a 0.1-0.2 mm needle into the heart without penetrating too deep into the ventricle (Figure 2B and Figure 3B).
   NOTE: If the tip end of the needle fills with blood, retract the needle slightly to avoid injecting the MS and QD into the ventricular lumen.
9. With the needle inserted into the heart apex, slightly raise the ventricle away from the body by reducing the angle between the needle and the body axis while moving the needle tip toward the base of the heart (Figure 2C and Figure 3C).
   NOTE: The needle may be faintly visible through the myocardial wall.
10. Push the needle further toward the head so the needle tip moves toward the myocardium surface.
11. Inject by pressing the pedal of the microinjection device for ~1-5 s or until a white spot is visible within the heart tissue (Figures 2C and Figure 3C,D).
12. If the chest cavity starts filling with injection fluid while injecting, it indicates that the needle tip has completely penetrated the myocardium (Figure 2D). Withdraw the needle slowly until a buildup of microbeads is visible within the tissue.
13. Reposition the needle tip if the fluid is injected into the ventricular lumen and cleared with the subsequent heartbeat (Figure 2E). In such a situation, lift the tip of the needle and push the needle toward the direction of the head until the injection pressure results in a white spot in the tissue.
14. If no white fluid or spot is visible, then the injection needle is blocked. In such a situation, retract the needle fully. To continue the experiment, either slightly break the tip of the needle or replace the needle. In both cases, check the injection flow prior to reinjecting the myocardium.
15. After successful injection, gently withdraw the needle from the myocardium (Figure 3F) and immediately transfer the zebrafish into a recovery tank with fish facility water.
16. Monitor the fish until it totally recovers from the tricaine.
17. Transfer the recovered fishes into a 2.8 L tank and place it in the fish facility, until the desired time point for heart extraction.

4. Heart extraction and imaging

1. Prepare the working concentration of tricaine by diluting 4.2 mL of stock solution (4 mg/mL) into 100 mL of fish facility water in a crystallizing dish.
2. Terminally anesthetize the zebrafish in tricaine solution until the movement of the gills has stopped and it is unresponsive.
3. Transfer the zebrafish to a moistened grooved sponge under the dissecting microscope positioned ventral side facing the objective lens.
4. Open the ventral wall of the chest with iridectomy scissors at the level of the pectoral fins.
5. Open the pericardial sac and locate the outflow track of the heart. Use forceps to grasp the aorta anterior to the Bulbous Arteriosus (BA) and carefully pull them up with the base of the ventricle.
6. With the anterior pole of the heart raised, cut the ventral aorta/sinus venosus to release the heart and place in a 35 mm Petri dish containing PBS.
7. Using forceps, remove any blood clots and fix the heart in 4% PFA (wt/vol) for 15 min.
8. Image the hearts using a microscope capable of acquiring the fluorescent wavelengths used.
   NOTE: In the example date, a confocal microscope with 405, 488, 567, and 647 laser lines was used.
9. Use ImageJ/Fiji software (see Table of Materials) for image analysis.
   NOTE: Calculation of QD or MS enrichment was performed using the high or low thresholding and create selection functions with the lymphatic vessel channel to first select regions of lymphatic vessels and those lacking vessel signal. Then, the Measure function was used to measure pixel intensity of the QD or MS channel from which a ratio of the two selections were calculated.

Results

Immediately after injection, a small white region of the myocardial wall must be visible (Figure 3F). This region will show bright fluorescent labeling of the injected MS and QD (Figure 4B,E). In addition, there may be weak and sporadic fluorescence puncta on the heart's outer surface from any QD and MS in the pericardial space following the procedure (Figure 4B,E). The injected tracers can be t...

Discussion

The present article has described a method to introduce exogenous material into the myocardium of zebrafish. This technique was developed to introduce QD and MS into the myocardium to study lymphatic function in homeostasis and regeneration^2,18. A similar approach has also been used to introduce QD into the myocardium of mice to investigate the presence and function of lymphatics after myocardial infarction^19,2...

Materials

Name	Company	Catalog Number	Comments
Crystallization dish	VWR	89000-288
Dissection Scope	Zeiss	495010-0007-000
Fish facility water	N/A	N/A	RO water with sea salt and sodium bicarbonate added to a conductivity of 226uS and pH of 7.35
Forceps	Dumont	11252-20
Glass Capillaries	WPI	1B120-3	no filament
ImageJ			https://imagej.nih.gov/ij/download.html
Iridectomy scissors	Fine Scientific Tools	15000-00
Microinjector	Warner Instruments	64-1735
Microloader femtotips	Eppendorf	5242 956.003
Micropipette puller	Sutter Instrument	P-97	Gated pedal input
Microspheres	Thermo Fisher Scientific	B200	Blue
PBS	Corning	46-013-CM
Quantum dots (QD)	Thermo Fisher Scientific	Q21061MP	Qtracker705 vascular label
Sponge	any	any	(1.5 × 5 × 3 cm) with groove (0.5 × 2.5 cm)
Syringe filter	Corning	431220
Tricaine	Sigma-Aldrich	A5040	concentration: 4 mg/mL