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 JoVE Biology

Intraperitoneal Injection into Adult Zebrafish

1, 2, 2,3, 1

1Department of Organismal Biology and Anatomy, The University of Chicago, 2Committee on Molecular Metabolism and Nutrition, The University of Chicago, 3Department of Medicine, The University of Chicago

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    Summary

    We demonstrate intraperitoneal injection into adult zebrafish. We use a 10 μl NanoFil microsyringe controlled by a Micro4 controller and UltraMicroPump III. This demonstration includes the use of cold water as an anesthetic.

    Date Published: 8/30/2010, Issue 42; doi: 10.3791/2126

    Cite this Article

    Kinkel, M. D., Eames, S. C., Philipson, L. H., Prince, V. E. Intraperitoneal Injection into Adult Zebrafish. J. Vis. Exp. (42), e2126, doi:10.3791/2126 (2010).

    Abstract

    A convenient method for chemically treating zebrafish is to introduce the reagent into the tank water, where it will be taken up by the fish. However, this method makes it difficult to know how much reagent is absorbed or taken up per fish. Some experimental questions, particularly those related to metabolic studies, may be better addressed by delivering a defined quantity to each fish, based on weight. Here we present a method for intraperitoneal (IP) injection into adult zebrafish. Injection is into the abdominal cavity, posterior to the pelvic girdle. This procedure is adapted from veterinary methods used for larger fish. It is safe, as we have observed zero mortality. Additionally, we have seen bleeding at the injection site in only 5 out of 127 injections, and in each of those cases the bleeding was brief, lasting several seconds, and the quantity of blood lost was small. Success with this procedure requires gentle handling of the fish through several steps including fasting, weighing, anesthetizing, injection, and recovery. Precautions are required to minimize stress throughout the procedure. Our precautions include using a small injection volume and a 35G needle. We use Cortland salt solution as the vehicle, which is osmotically balanced for freshwater fish. Aeration of the gills is maintained during the injection procedure by first bringing the fish into a surgical plane of anesthesia, which allows slow operculum movements, and second, by holding the fish in a trough within a water-saturated sponge during the injection itself. We demonstrate the utility of IP injection by injecting glucose and monitoring the rise in blood glucose level and its subsequent return to normal. As stress is known to increase blood glucose in teleost fish, we compare blood glucose levels in vehicle-injected and non-injected adults and show that the procedure does not cause a significant rise in blood glucose.

    Protocol

    1. Pre-injection Preparations

    1. Fast the fish for at least 24 hours prior to injection. This will empty the intestinal bulb (stomach) contents. The basic fasting protocol is to transfer the fish, at their normal density, to a clean tank, then withhold food. For longer-term fasting that requires more rigorous conditions (e.g., for blood glucose studies), see additional considerations in the Discussion.
    2. Prepare Cortland salt solution (Perry et al., 1984).
      For a 100 mL volume, dissolve the following in distilled water:
      725 mg NaCl (124.1 mM)
      38 mg KCl (5.1 mM)
      41 mg Na2HPO4 (2.9 mM)
      24 mg MgSO4∙7H2O (1.9 mM)
      16 mg CaCl2∙2H2O (1.4 mM)
      100 mg NaHCO3 (11.9 mM)
      4 g Polyvinylpyrrolidone (PVP) (4%)
      1,000 USP units Heparin
      Filter, sterilize and store at 4°C.
    3. Prepare the microscope.
      • Cover the microscope base with plastic wrap for protection in case of spills.
      • Put a paper towel on top of the plastic wrap. The surgical table will sit on top of the paper towel.
      • Pre-adjust focus by viewing the surgical table and focusing on the sponge.
      Tip: Put your finger on top of the sponge and focus on that. This will eliminate or minimize further focal adjustment once the fish is on the surgical table.
    4. Weigh the fish.
      • Fill a 500 mL beaker about 1/3 full with fish facility water.
      • Tare the balance.
      • Collect the fish using a net. Wick excess water away from the net and fish by briefly dabbing the net on paper towels. Transfer the fish to the beaker.
      • Weigh the fish.
      • Transfer the fish to a clean tank.
      • Transfer each weighed fish to its own labeled tank.
      • Calculate the injection volume for each fish based on fish weight.
    5. Prepare the syringe and related injection equipment. For injection, we recommend a 35G beveled steel needle and a 10 μl NanoFil microsyringe. Prepare the NanoFil syringe and silflex tubing following the manufacturer's instructions. It is important to remove any bubbles from the syringe and tubing. After filling the syringe and tubing, mount the syringe on the pump, and program the injection volume for the first fish.
    6. Prepare the surgical table.
      • Cut a soft sponge (such as #L800-D, Jaece Industries) so that it is approximately 20 mm in height. On the flat face, make a cut that is 10-15 mm deep. This cut is the trough that will hold the fish for injection.
      • Set the sponge into a 60 mm Petri dish.
      • Set the Petri dish with sponge into a suitably-sized pipette tip box lid. The lid needs to be large enough to hold water to help maintain sponge temperature, but it should be shallow enough to not get in the way. We use a lid from a P200 tip box that is 11.4 cm L x 7.7 cm W x 1.5 cm D.
      These three items assembled together (sponge in petri dish in box lid) constitute the surgical table.
    7. Prepare the anesthetic.
      • Make crushed ice using cubes made from fish facility water.
      Tip: Using typical ice cube trays, it will take 3 trays to anesthetize 10-12 fish.
      • Fill a clean ice bucket with the crushed ice.
      • Put the surgical table into a larger container such as a 2.4 liter Rubbermaid food storage container.
      • Pour some facility water (warm) into the outer container and the surgical table. Keep a reserve of warm facility water nearby.
      • Put a thermometer into the outer container.

    2. Anesthesia, Injection and Recovery

    1. Place the anesthetic outer container plus surgical table adjacent to the microscope. Have the bucket of ice chips nearby.
    2. Bring the water temperature down to 17°C by adding ice chips. Important: Don't go below 17°C for this step.
    3. Use a net to transfer the fish to the outer container.
    4. Slowly add ice chips to the container to bring the temperature down to 12°C, over the course of several minutes.
    5. Monitor fish behavior: At 17°C or slightly lower, the fish typically will spread its pectoral fins horizontally, gasp, and have rapid operculum movements. As the temperature drops, the fish will swim more slowly and finally stop swimming. As the surgical plane of anesthesia is approached, gasping will stop and operculum movements will slow. The fish is ready for injection when it does not react to being handled. For most fish, 12°C is sufficient. Larger fish may require colder water.
    6. As the required temperature is reached (~12°C or colder), press on the sponge to saturate it.
    7. Keep your fingers in the cold water sufficiently so that they will not warm up the fish and bring it out of anesthesia during handling.
    8. With cold fingers, gently transfer the fish to the trough of the sponge. Position the fish with the abdomen up and the gills in the trough.
    9. Quickly transfer the surgical table to the microscope stage.
    10. Working quickly, carefully insert the needle into the midline between the pelvic fins. The needle should point cranially and be inserted closer to the pelvic girdle than to the anus. You should be able to feel when the needle is deep to the body wall. Inject the appropriate volume and withdraw the needle.
    11. After injection, immediately transfer the fish back to its warm-water (~28.5°C) tank for recovery by releasing the fish from the sponge over the tank water.
      Tip: If the fish does not begin swimming immediately, help it to recover by gently swirling water towards the gills.
    12. Check the needle. Occasionally a scale may be attached and should be removed prior to the next injection.
    13. For subsequent injections, use warm facility water to bring the anesthetic chamber water temperature back up to 17°C before introducing the next fish.

    3. Representative Results:

    Figure 1
    Figure 1. Representative results following intraperitoneal injection of 0.5 mg/g glucose or vehicle. Fish were fasted for 72 hours prior to injection. The x-axis shows time, post-injection. Mean ± SEM.

    Discussion

    Intraperitoneal injection involves five steps: fasting, weighing, anesthetizing, injection, and recovery. For each step there are best practices that can ensure success. Success includes a healthy fish patient as well as a good experimental outcome.

    Fasting: A 24-hour fast should empty the intestinal bulb. This practice is taken from the fish veterinary literature (e.g., Brown 1993). Additional fasting considerations are discussed below.

    Longer-term fasting: We have found that a 72-hour fast is required to decrease blood glucose to a baseline level prior to injection (Eames et al., 2010). We have also found that for glucose studies there are several procedures that are required to ensure that the fish are fasted properly. Start with a clean tank (no debris on the bottom). Tanks should be offline, clearly labeled as 'fasting', and in a location where enthusiastic fish care personnel will not feed them. Evaluate the external environment of the tank and take steps to prevent the fish from being stressed from disturbances, as stress is known to raise blood glucose (Chavin and Young, 1970; Groff et al., 1999). For example, we had a fasting experiment in which a radio was operated daily on the bench that was holding the fish tanks. We found that blood glucose was unusually high and concluded that the fish were stressed by the vibrations. Another stressor is overcrowding. Fish should be kept at a density that conforms with good fish husbandry practices. For recommendations, see Brand et al. (2002) and Westerfield (1995). We have had good results fasting our fish at a density of 10-12 fish in a 9 liter tank (with 3 layers of marbles taking up some of that volume). Separating the sexes may cause stress, so we recommend maintaining a mixed-sex population during the fast. This means that eggs can be laid, and the eggs need to be sequestered so that they will not be eaten. A simple way to sequester eggs is to cover the tank bottom with 2-3 layers of marbles. Water quality needs to be maintained by removing eggs and waste and by replacing about 10-15% of the tank water, daily. For removing eggs and waste, siphoning works well.

    Weighing: When weighing fish that are not anesthetized, care should be taken to minimize water transfer from the net into the beaker, to ensure accurate weighing. If the net (with fish) is blotted on paper towels, the majority of the excess water can be removed, and the weight can be accurately measured. It may be easier to anesthetize the fish prior to weighing, but we have not tested the possible effects of anesthetizing a fish twice in one day. We have tested our technique by weighing the fish first with the netting/blotting method and then re-weighing the fish after it has been anesthetized, and gently blotted dry. We found no significant difference in weight between the methods (P = 0.7927, t-test). Additionally, we tested whether this netting/blotting method affected blood glucose, in comparison with simply transferring the fish to the beaker as soon as it is netted (no blotting). We found no significant difference in blood glucose level between the two transfer methods (P = 0.2241, t-test).

    Anesthetizing: Chemical anesthesia may be suitable for many studies. Here we have demonstrated cold water anesthesia as an alternative, because many anesthetics (including tricaine/MS-222 (Brown, 1993)), raise blood glucose. In previous studies, we have determined that cold water does not raise blood glucose in zebrafish (Eames et al., 2010).
    For cold water anesthesia, the temperature should be decreased slowly. The rate of decrease seems to depend on the size of the fish, with smaller fish going under faster than larger fish. Following injection, you may observe that the fish is recovering too slowly from the anesthetic (see below). This can result when either the starting temperature is too low, or when the temperature is decreased too rapidly. The starting temperature is too low if the fish bends laterally upon entering the water. If the starting temperature is correct, the fish will maintain its balance initially. It will rotate its pectoral fins to a horizontal position, gasp, and have rapid operculum movements. Typically, it will swim. As temperature decreases, movements will decrease and the fish will lose equilibrium. A surgical plane of anesthesia is reached when the fish can be handled without reacting. To maintain the fish under surgical anesthesia, your fingers must be cold, so keep them in the water prior to handling the fish. The sponge must also be kept cold at the same temperature as the water used for anesthetizing the fish. It is important to saturate the sponge with water that is sufficiently cold to maintain anesthesia once the fish is placed onto it.

    Injection: Prior to undertaking injections, you may want to dissect at least one fish to get a sense of body wall thickness. This can help you to judge how far the needle needs to insert to enter the abdominal cavity. Additionally, as you insert the needle, you can feel the body wall "give" when the needle enters the abdominal cavity. During the injection, take steps to keep the patient happy. Make sure the sponge is saturated with the correct temperature cold water to prevent the fish from reviving during injection. A well-saturated and soft sponge is important for minimizing damage to the scales and mucus covering of the skin. A well-saturated sponge is also important for keeping the gills aerated. We highly recommend the foam sponge listed below under Materials. Finally, once the fish is anesthetized, work quickly to minimize the time that the fish is under.

    Recovery: The fish should recover from the anesthesia virtually upon entering the warm tank water. If the fish does not begin swimming immediately, gently swirl the water towards its gills to speed recovery. If recovery is slow, then the fish went under too quickly and you should adjust the anesthesia procedure appropriately. The possible causes of slow recovery are discussed under Anesthetizing.

    Disclosures

    No conflicts of interest declared.

    Acknowledgements

    This study was supported by Juvenile Diabetes Research Foundation grant 5-2007-97 (to V.E.P.), by National Institute of Diabetes and Digestive and Kidney Diseases grants R01DK064973 (to V.E.P.), R01DK48494 (to L.H.P.), T32DK07074 (supporting S.C.E.), K01DK083552 (to M.D.K), and by P60DK20595 to The University of Chicago Diabetes Research and Training Center. The content is solely the responsibility of the authors and does not necessarily represent the official views of the NIDDK or the NIH.

    Materials

    Name Company Catalog Number Comments
    Foam Sponge Jaece Industries L800-D
    60 mm Petri dish
    Pipet tip box lid not too deep, e.g. 1.5 cm
    Plastic storage container deep, e.g. 7 cm
    Thermometer
    Crushed ice made from facility water
    Warm facility water 1 liter or more
    500 ml beaker for weighing
    NanoFil syringe World Precision Instruments, Inc. NANOFIL or Hamilton syringe
    35 gauge needle World Precision Instruments, Inc. NF35BV-2 beveled
    Silflex tubing World Precision Instruments, Inc. SILFLEX-2
    UltraMicroPump III and Micro4 controller World Precision Instruments, Inc. UMPS-1
    Foot switch World Precision Instruments, Inc. 15867
    Dissecting microscope
    Plastic wrap
    Paper towels
    Cortland salt solution

    References

    1. Perry, S.F., Davie, P.S., Daxboeck, C., Ellis, A.G. & Smith, D.G. Perfusion methods for the study of gill physiology. In Fish Physiology Volume X: Gills, Part B: Ion and Water Transfer. Hoar, W.S. & Randall, D.J. (eds), pp. 325-388, Academic Press, Inc., Orlando (1984).
    2. Brown, L.A. Anesthesia and restraint. In Fish Medicine. Stoskopf, M.K. (ed), pp. 79-90, W.B. Saunders Company, Philadelphia (1993).
    3. Eames, S.C., Philipson, L., Prince, V.E. & Kinkel, M.D. Blood sugar measurement in zebrafish reveals dynamics of glucose homeostastis. Zebrafish 7, 205-213 (2010)..
    4. Chavin, W. & Young, J.E. Factors in the determination of normal serum glucose levels of goldfish, Carassius auratus L. Comp Biochem Physiol 33, 629-653 (1970).
    5. Groff, J.M. & Zinkl, J.G. Hematology and clinical chemistry of cyprinid fish. Common carp and goldfish. Vet Clin North Am Exot Anim Pract 2, 741-776 (1999).
    6. Brand, M., Granato, M. & Nusslein-Volhard, C. Keeping and raising zebrafish. In Zebrafish: A Practical Approach. Nusslein-Volhard, C. & Dahm, R. (eds), pp. 7-37, Oxford University Press, Oxford (2002).
    7. Westerfield, M. The Zebrafish Book University of Oregon Press, Eugene (1995).
    8. Iwama, G.K. & Ackerman, P.A. Anaesthetics. In Biochemistry and Molecular Biology of Fishes, Volume 3: Analytical Techniques. Hochachka, P.W. & Mommsen, T.P. (eds), pp. 1-15, Elsevier, Amsterdam (1994).
    9. Reavill, D.R. Common diagnostic and clinical techniques for fish. Vet Clin North Am Exot Anim Pract 9, 223-235 (2006).
    10. Stoskopf, M.K. Surgery. In Fish Medicine. Stoskopf, M.K. (ed), pp. 91-97, W.B. Saunders Company, Philadelphia (1993).

    Comments

    22 Comments

    Hi, I m an animal technician and thi article is very good for me. Thanks for this work.
    Carolina Mourelle (Buenos Aires, Agentina)
    Reply

    Posted by: AnonymousSeptember 3, 2010, 5:54 PM

    how can i measure blood glucose of zebrafish?which process do you offer me?thanks
    Reply

    Posted by: AnonymousDecember 5, 2010, 2:10 AM

    Hi,
    Detailed methods for blood collection and measuring blood glucose in zebrafish are in reference #3 above. In that paper we showed that both the OneTouch meter and the FreeStyle meter are accurate with zebrafish blood. In practice, we prefer the FreeStyle meter because it uses a smaller blood volume and is faster than the OneTouch. However, the test strip chemistry for the FreeStyle meter was recently changed by the manufacturer, and we are in the process of testing whether the new strips perform as well as the old strips.
    Mary
    Reply

    Posted by: AnonymousDecember 5, 2010, 11:44 AM

    Thanks for the very helpful video! I am hoping to start measuring blood glucose in zebrafish, and I would like to use the FreeStyle meter. Have you done any more testing on the new FreeStyle test strips? I would love to know if these are as effective and consistent as the old test strips.
    Thanks for your help, and thanks again for all your work!
    Reply

    Posted by: AnonymousDecember 16, 2011, 6:27 PM

    Great video. Thank you for posting.
    Reply

    Posted by: AnonymousMay 5, 2011, 4:11 PM

    Thanks!
    Mary & Stef
    Reply

    Posted by: Mary K.May 5, 2011, 5:01 PM

    Really very nice and much needed information.congratulations.I am Kirty Sirothia,Associate Prof.Vet Path.Nagpur Veterinary College,India.This type of basic work video is very necessary for furthering the science.
    Reply

    Posted by: AnonymousMay 8, 2011, 8:59 AM

    Thanks!
    Stef & Mary
    Reply

    Posted by: Mary K.May 9, 2011, 11:15 AM

    What is the limit of injection volume for average size zebrafish (0.3~0.5g)?
    Thank you!
    Reply

    Posted by: AnonymousAugust 8, 2011, 3:42 AM

    Very helpful. Thanks from Vancouver, BC.
    Reply

    Posted by: AnonymousAugust 16, 2011, 3:23 PM

    I work in an aquatic toxicology laboratory at The University of Massachusetts Amherst. The model organism we use is Japanese Medaka. We are currently developing a method to inject gold nano particles into Medaka to see if they cross the blood brain barrier, which could be beneficial to drug delivery. Your video was very helpful in preparing the methodology. We would now like to go further and perform an experiment. I was wondering if it would be possible for you to share your IACUC protocol with our lab. It would be greatly appreciated. Thank you so much for publishing this video.
    Best,
    Kasie Auger
    Reply

    Posted by: AnonymousNovember 22, 2011, 4:54 PM

    Just wanted to suggest a small but significant change to the protocol shown here. Instead of using cold water to anesthetize zebrafish (which can be extremely painful), use 80-90 mg/L MS-²²² (Tricaine). This is the standard and ethical way to anesthetize fish. Euthanasia can also be acheived at higher doses.
    Reply

    Posted by: AnonymousNovember 22, 2011, 5:07 PM

    Hi Jeremy,
    We appreciate your comments about anesthesia choice and take your concerns very seriously. When we were developing these methods, we did extensive reading of the fish veterinary literature and found that cold water anesthesia for teleost fish is a standard method (although not the laboratory standard for zebrafish, as you point
    out). Because cold water anesthesia is an established method, we have not previously had anyone suggest that there may be an ethical issue with its use. We consider the ethical care of zebrafish to be extremely important.
    We know from the literature and from colleagues that IACUCs at multiple institutions approve the use of cold
    water for anesthetizing zebrafish. In fact, cold water is also acceptable for euthanizing zebrafish, and an example
    is the IACUC protocol approved for ZIRC, which can be viewed on the ZIRC website. The main differences with anesthetizing versus euthanizing is that with anesthetizing the water temperature is gradually lowered rather than introducing an abrupt change, and the temperature gŒs down to typically 1² degrees Celsius, rather than the
    much icier water required for euthanizing. All procedures shown in the video were approved by the Institutional Animal Care and Use Committee at The University of Chicago. This statement is included in the video version of
    the protocol starting at 15 seconds from the beginning.

    We have read the literature on teleost pain sensation, and have not found evidence that lowering the water temperature to 17-1² degrees Celsius would induce pain. Natural history studies have shown that zebrafish
    tolerate a wide temperature range in their native habitats. For example, Spence et al. ²008 reported that winter
    water temperature on the Indian subcontinent dropped as low as 6 degrees Celsius. However, such studies have
    not measured the potential effect of temperature on pain sensation. Can you help us out with references for this?

    Reply

    Posted by: Mary K.November 30, 2011, 6:56 PM

    ...just read your note about cold water anesthasia (above). Obviously for the rest of us who aren't measuring blood glucose levels, Tricaine should be the preferred option. Cheers.
    Reply

    Posted by: AnonymousNovember 22, 2011, 5:12 PM

    First of all, I appreciate your concern for fish welfare and by no means do I feel holier than thou regarding this topic. Secondly, I appreciate this video demonstration and have followed it myself. Finally, thanks for your reply to my post.

    Re. pain sensation in fish, I can't say for certain the gradual reduction of water temperature causes pain in fish (my previous assertion was meant to communicate the possibility of pain - I incorrectly used the word "can", and should have used the word "may"). I am cynical about any procedure that operates under the presumption of no pain. Although wild zebrafish do tolerate a range of temperatures, they probably do not experience temperatures cold enough to knock them out (assumption).

    At any rate, I do recognize that cold water anesthesia is acceptable in some institutions and I'm not going to debate their choice to follow that protocol. However, if given the option, I believe erring on the side of caution is the way to go.
    Reply

    Posted by: AnonymousDecember 1, 2011, 2:06 PM

    What is the maximum volume one can inject an adult zebrafish? 10ul? more? less?
    Reply

    Posted by: Irene B.March 14, 2013, 10:32 AM

    Hi Irene, Thanks for your question. We need to look back in our notes and we'll get back to you soon.
    Reply

    Posted by: Stefani (.March 14, 2013, 2:41 PM

    Hi Irene,
    Although I gave you a longer answer by email, I figured I'd post a short answer here in case other people have the same question. We did not test the maximum volume that could be injected, so I don't have a good answer. All we can say is that, in theory, a larger fish should be able to tolerate a larger injection volume than a smaller fish. The largest volume we injected was about 1.² microliters and the fish seemed to tolerate it fine. However, our typical injection volume was between about ²50 and 450 nanoliters.
    Mary
    Reply

    Posted by: Mary K.April 4, 2013, 9:43 AM

    Hi, thank you for publishing this procedure. I am injecting zebrafish intramuscularly (just starting to practice the technique), and found a lot of helpful tipse in your video! Could I ask you about the specifications for the microscope you use? Best , Janicke Nordgreen.
    Reply

    Posted by: Janicke N.April 4, 2013, 3:06 AM

    Hi Janicke,
    Thanks for the comments. We used a Leica M165 FC. The features you need in a scope are a wide base to help you steady your hands, and a good working distance. Good luck!
    Mary
    Reply

    Posted by: Mary K.April 4, 2013, 9:30 AM

    Hey,

    I was just wondering what method you used to extract blood? decapitation? How many microlitres did you manage to collect?

    Best,

    Andrew

    Reply

    Posted by: Andrew L.January 9, 2014, 10:19 AM

    Hi Andrew,
    Please refer to reference 3 above, in which we published detailed methods for blood collection following decapitation. We were usually able to collect approximately 5 ul of blood but easily could get up to 10 ul from larger fish.
    Good luck with your work,
    Stef
    Reply

    Posted by: Stefani (.January 16, 2014, 6:19 PM

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