Repeated Blood Collection for Blood Tests in Adult Zebrafish

Summary

Repeated blood sampling is necessary and important in animal research. We developed a novel, non-lethal and reliable method for repeated blood collection from adult zebrafish, and applied this method to the study of blood biochemistry, including glucose metabolism.

Abstract

Repeated blood collection is one of the most common techniques performed on laboratory animals. However, a non-lethal protocol for blood collection from zebrafish has not been established. The previous methods for blood collection from zebrafish are lethal, such as lateral incision, decapitation and tail ablation. Thus we have developed a novel “repeated” blood collection method, and present here a detailed protocol outlining this procedure. This method is minimally invasive and results in a very low mortality rate (2.3%) for zebrafish, thus enabling repeated blood sampling from the same individual. The maximum volume of blood sampling is dependent on body weight of the fish. The volume for repeated blood sampling at intervals should be ≤0.4% of body weight every week or ≤1% every 2 weeks, which were evaluated by measurements of blood hemoglobin. Additionally, hemoglobin, fasting blood glucose, plasma triacylglycerol (TG) and total cholesterol levels in male and female adult zebrafish were measured. We also applied this method to investigate the dysregulation of glucose metabolism in diet-induced obesity. This blood collection method will allow many applications, including glucose and lipid metabolism and hematological studies, which will increase the use of zebrafish as a human disease model organism.

Introduction

Zebrafish are gaining increasing popularity as a valuable model of human diseases because their organs and genetics are similar to those of humans^1,2. In the field of developmental biology, many studies have demonstrated that zebrafish and human show marked similarity in hematopoiesis³, hemostasis^4,5, and myelopoiesis⁶. Adult zebrafish are also used for studying immunological⁷, neurodegenerative⁸ and obesity-related diseases⁹ because this model organism shares common pathways with those disrupted in human diseases. For obesity and obesity-related diseases (diabetes, hepatic steatosis and nonalcoholic steatohepatitis and atherosclerosis), zebrafish blood glucose and lipids levels have been thoroughly investigated in several transgenic and diet-induced obesity models^10-13.

Repeated blood sampling from individual animals will reduce animal use and decrease interindividual differences. However, repeated sample collection is technically difficult in small animals such as zebrafish because of their relatively small blood volume and the lack of easily accessible vessels. Several methods for one-time blood collection from zebrafish have been developed, although these methods have their own drawbacks, including lethality, associated tissue damage and limited blood volume. For example, 1–5 µl blood can be harvested from a lateral incision of approximately 0.3 cm in length in the region of the dorsal aorta⁵. Decapitation with scissors by cutting through the pectoral girdle can collect 5–10 µl blood¹⁰. Another convenient blood sampling method is tail ablation¹⁴. Cardiac puncture is one potential alternative method for repeated blood collection from the same fish, but the very small amount obtained (approximately 50 nl) with this procedure limits the number of analyses that can be performed¹¹. Accordingly, a new protocol is needed to enable repeated non-lethal blood sampling, which would be a critical advance necessary for this organism to be a standard model organism for human diseases. This technique would allow for testing pharmacological response, discovery of molecular biomarkers for diagnosis, determination of prognosis, and the monitoring of various diseases, such as metabolic diseases, degenerative diseases and several types of malignancies.

We therefore developed a minimally invasive method for obtaining blood from zebrafish serially¹⁵. Here we demonstrate the procedure visually and provide a detailed protocol for this technique. Using this method, the normal value based on various parameters, including hemoglobin, fasting blood glucose, and lipids in the blood of healthy adult zebrafish were evaluated. Additionally, we also evaluated whether this method is suitable for studies that require serial samples by monitoring the temporal changes in blood glucose levels during overfeeding experiments.

Protocol

All animal procedures were approved by the Ethics Committee of Mie University, and were performed according to Japanese animal welfare regulation ‘Act on Welfare and Management of Animals’ (Ministry of Environment of Japan) and complied with international guidelines.

1. Preparation of the Needle  

NOTE: All experiments were performed under anesthesia, and all efforts were made to minimize suffering. For euthanasia, fish were immersed in an ice–water bath (5 parts ice/1 part water at ≤4 °C) for ≥20 min.

1. Prepare the glass microcapillary needles by pulling a 1.0-mm-outer-diameter glass capillary with a needle puller (Figure 1A).
2. Cut the tips of the needles obliquely using fine scissors (Figure 1B). The ideal tip diameter should be approximately 100 – 200 µm (Figure 1C). If the tip diameter is too narrow, blood will not enter the needle.
3. Dissolve heparin in saline to a concentration of 5 mg/ml.
4. Place a precut needle in the nosepiece end of an aspirator tube assembly and hold the mouthpiece in the mouth, or connect the needle to a bulb dispenser. Immerse the needle tip into the heparin solution, and heparinize the needle by suction and blowing through the solution (Figure 1D and 1E).
   Note: The aspirator tube assembly has been used for zebrafish sperm cryopreservation¹⁶. Long rubber tube can stop any blood flying into mouth. Setting a filter in the midway of the tube can avoid the hazards.
5. Store the heparinized needles in a 10-cm Petri dish and air-dry for at least 1 hr. A large number of needles can be prepared in advance.

2. Anesthesia

1. Prepare the anesthetic solution in a small plastic case by mixing 200 ml of fish water with 100 µl of 2-phenoxyethanol (2-PE). Final concentration of the anesthetics is 500 ppm.
   CAUTION! 2-PE has a rapid effect.
2. Remove desired number of fish (AB strain) from the circulation system.
3. Use a net to transfer the fish into the anesthetics for 1 – 2 min (Figure 1F). Observe the fish gradually swim, spread the pectoral fins horizontally, gasp, and have rapid operculum movements within 1 min.
4. As the time goes on, observe the fish lay on the bottom of the case and finally stop swimming. The surgical plane of anesthesia will reach when the fish stops to gasp and the operculum movements are slow. At this point, fish is ready for blood collection.
5. Using a skimmer lift the anesthetized fish from the 2-PE and gently place it on a paper towel soaked with the anesthetics ( Figure 1G). Cover the fish’s head with soft tissue paper also soaked with 2-PE solution to prevent eye dryness and use another dry soft tissue paper to gently dry off the body surface.

3. Blood Collection

1. Place a heparinized needle in the nosepiece end of the aspirator tube assembly (or the bulb dispenser) and hold the mouthpiece end of the aspirator tube assembly in mouth.
2. Grasp the nosepiece end and the needle together, and carefully remove the interfering scales with the needle tip. Insert the needle at a 30 – 45° angle into the blood collection site. Avoid puncture of the gastrointestinal tract (Figure 1H). In case of using bulb dispenser, press the bulb with thumb and middle finger, and block the hole at the tip of the bulb with first finger, then insert the needle as described above.
   Note: The site for blood collection is along the body axis and posterior to the anus in the region of the dorsal aorta. The dorsal aorta (DA) and the posterior cardinal vein (PCV) are just ventral to the spine (Figure 2).
3. Begin to suck the mouthpiece end of the aspiration tube assembly when the needle is felt touching the spine. If blood does not rise, move the needle tip subtly by hand to encourage blood flow. Note that once blood is rising into the needle, immediately stop shaking and suck gently (Figure 1I). In case of using bulb dispenser, release the pressure of bulb to aspirate the blood.
4. Observe the blood will slowly rise into the needle in a pulsatile manner without suction, which is likely because of the arterial blood pressure. Thus, it is not necessary to suck if the needle correctly penetrates the artery.
5. Stop suction after the appropriate volume of blood is collected. Remove the needle from the fish and press the puncture site using soft tissue paper to stop any bleeding. Observe the bleeding stop after approximately 10 – 20 sec of finger pressure (Figure 1J).
6. After the bleeding is stopped, immediately transfer the fish back to a clean warm–water (~ 28 °C) tank. Help the fish to recover by gently swirling water towards the gills until it begins to swim.
   Note: Keep up to 5 postsampling fish in a 2 L tank and connect the tank to the circulation system to oxygenate the fish. Addition of antibiotics to the fish water is not necessary. Maintain the fish with normal housing and feeding.
7. Expel the blood from the needle onto a clean area of a piece of parafilm (Figure 1K).
8. Measure blood glucose using any commercial handheld glucometer (Figure 1L).
   Note: The glucometer uses a glucose dehydrogenase–flavin adenine dinucleotide electrode and requires a sample volume of 0.6 µl.
   1. Insert a test strip completely into the meter and directly touch the blood drop. Observe the blood draw into the test strip automatically and obtain the blood glucose result 5 sec on the display area. Record the result and discard the test strip. Use a new test strip for each measurement.
9. (optional step) Take accurate quantities of blood by a pipetting and transfer the blood to a microcentrifuge tube for further analyses (hemoglobin, triacylglycerol (TG), total cholesterol, etc.). If necessary, dilute the whole blood from one fish with saline. Centrifuge the blood samples for 3 min at 680 x g at RT and harvest the plasma. Transfer the plasma into a new tube. At this point, it is ready to be used in biochemical analyses.

Results

This blood collection method causes minimal injury to zebrafish (a <1 mm puncture; Figure 1J) and yields a very low mortality rate of 2.3%. We examined the maximum volume of blood that could be collected from a single fish and evaluated the relationship to its body weight (Figure 3). We found that the maximum blood volume collected was linearly correlated with body weight (R = 0.813). The largest volume of blood collected from an individual fish (body weight = 1.071 g) was 25 µl, and ...

Discussion

We present here a detailed protocol for serially obtaining blood from adult zebrafish. This method is straightforward to carry out and we use it in lab on a daily basis. This blood collection method is based on inserting a glass capillary needle into the zebrafish’s dorsal aorta. During this procedure, it is critical to be careful to not ablate the spine because it is the criterion for searching for the dorsal aorta. Reducing the spine injury will improve the survival rate. Although this technique is simple and eas...

Materials

List of materials used in this article

Name	Company	Catalog Number	Comments
Glass capillaries with filament	Narishige	GD-1	1.0 mm outer diameter.
Needle puller	Narishige	PC-10	To produce the needles
Heparin	Wako Pure Chemical Industries	081-00136	For heparinization
Aspirator tube assembly	Drummond	2-040-000	For blood collection
Bulb dispenser	Drummond	1-000-9000	For blood collection
2-phenoxyethanol	Wako Pure Chemical Industries	163-12075	For anesthetizing the fish
DRI-CHEM3500V	Fujifilm	-	For hemoglobin measurement
DRI-CHEM Slides	Fujifilm	Hb-WII	For hemoglobin measurement
Glutest Neo Super	Sanwa Kagaku Kenkyusho	-	For bood glucose measurement
Wako L-type TG kit	Wako Pure Chemical Industries	464-44201	For TG measurement
Wako L-type CHO kit	Wako Pure Chemical Industries	460-44301	For total cholesterol measurement
Parafilm M	Alcan Packaging	PM996	To expel the blood on