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. Author manuscript; available in PMC: 2023 Oct 7.
Published in final edited form as: Methods Enzymol. 2018 Mar 15;602:189–209. doi: 10.1016/bs.mie.2018.02.004

Zebrafish: A Pharmacogenetic Model for Anesthesia

Victoria Bedell *,1, Elena Buglo §,¶,‖, Daniel Marcato †, Christian Pylatiuk ‡, Ralf Mikut ‡, Johannes Stegmaier ‡, Will Scudder ‖, Maxwell Wray ‖, Stephan Züchner §,, Uwe Strähle †, Ravindra Peravali †, Julia E Dallman ‖
PMCID: PMC10559369  NIHMSID: NIHMS1932043  PMID: 29588029

Abstract

General anesthetics are small molecules that interact with and effect the function of many different proteins to promote loss of consciousness, amnesia, and sometimes, analgesia. Owing to the complexity of this state transition and the transient nature of these drug/protein interactions, anesthetics can be difficult to study. The zebrafish is an emerging model for the discovery of both new genes required for the response to and side effects of anesthesia. Here we discuss the tools available to manipulate the zebrafish genome, including both genetic screens and genome engineering approaches. Additionally, there are various robust behavior assays available to study anesthetic and other drug responses. These assays are available for single-gene study or high throughput for genetic or drug discovery. Finally, we present a case study of using propofol as an anesthetic in the zebrafish. These techniques and protocols make the zebrafish a powerful model to study anesthetic mechanisms and drug discovery.

1. INTRODUCTION

Despite the widespread use of anesthetics for surgical practice, the contribution of genetics to variability in individual responses to anesthesia is still poorly understood (Garcia, Rothman, & Fitzpatrick, 2017; Mashour, Woodrum, & Avidan, 2015; Saczynski et al., 2012). For example studies of red-haired women, who anecdotally had required increased anesthetic, actually verified an association with increased anesthetic requirement over their dark haired matched counterparts (Liem et al., 2004). This illustrates a genetic component to anesthetic response that is not well understood. Additionally, classic, large-scale studies show that, irrespective of age, 3%–8% of people experience prolonged grogginess for more than 2h after surgery with unknown cause (compared to ~80% that recover within 15min) (Aldrete & Kroulik, 1970; Philip, Kallar, Bogetz, Scheller, & Wetchler, 1996). Postoperative cognitive issues are both more severe and more prevalent in patients over the age of 60, as up to 40% experience delirium associated with negative prognoses after undergoing inpatient surgeries (Giattino et al., 2017). Other complications include prolonged hospital stays as well as increased morbidity and mortality (Mashour et al., 2015; Neufeld et al., 2013; Saczynski et al., 2012). It has also been shown that reduced cognitive ability before anesthesia correlates with greater risk for cognitive decline postanesthesia (Giattino et al., 2017). To address the contributions of genetics and environment to postoperative delirium, a large twin study of middle aged and elderly patients strongly suggest that surgery/anesthesia alone are not sufficient to explain such complications (Dokkedal, Hansen, Rasmussen, Mengel-From, & Christensen, 2016; Terrando et al., 2011). Taken together these data support the possibility of an individual’s genetic make-up can profoundly impact in their response to anesthetics and that inherited neurological disorders contribute their perioperative experiences.

Since the sequencing of the human genome in 2001, thousands of genetic variants that cause inherited disorders have been identified (Huguet, Benabou, & Bourgeron, 2016; Puschmann, 2013; Rudnik-Schoneborn, Auer-Grumbach, & Senderek, 2017). Case studies link rare inherited nervous system disorders such as narcolepsy and glycine encephalopathy to complications after anesthesia that include cataplexy and trouble breathing (Liu & Fan, 2006; Mesa, Diaz, & Frosth, 2000). Perioperative risks are also well established for more common inherited disorders such as epilepsy and Parkinson’s disease (Dhallu, Baiomi, Biyyam, & Chilimuri, 2017). Therefore, it has been hypothesized that at least a portion of an individual’s sensitivity to anesthetics and postoperative complications could result from preclinical inherited neurological disorders (Dhallu et al., 2017; Dokkedal et al., 2016).

Zebrafish provide a promising emerging model in which to better understand genetic variants that confer risk with anesthesia. Established as a new model organism in the 1970s and 80s by George Streisinger at the University of Oregon (Streisinger, Walker, Dower, Knauber, & Singer, 1981), zebrafish (Brachydanio rerio) combine the advantages of two more established model organisms, the fruit fly and the mouse. Like fruit flies, a single zebrafish female produces hundreds of embryos at a time that develop externally and rapidly acquire a diversity of sensory-evoked locomotor behaviors (Orger & de Polavieja, 2017) that can be used to assess depth of sedation. In addition, embryos/larvae are transparent making it possible to track neuronal activity in vivo (Ahrens, Huang, Narayan, Mensh, & Engert, 2013) to determine neural circuit correlates of anesthesia. Like mice and humans, zebrafish are vertebrates with largely conserved genes (Howe et al., 2013) and similar physiology, including ion channels, neurotransmitters, and receptors (Renier et al., 2007; Rihel & Schier, 2013) which are known to be among the targets of anesthetic agents (Forman & Miller, 2011; Kretschmannova et al., 2013; Orser, Wang, Pennefather, & MacDonald, 1994). Combined with flexible methods for mutagenizing the genome that includes both forward (Lawson & Wolfe, 2011) and reverse genetic approaches (Hsu, Lander, & Zhang, 2014) and high-throughput methods for behavioral screening (Kokel & Peterson, 2011; McCarroll, Gendelev, Keiser, & Kokel, 2016; Rihel et al., 2010), we suggest that zebrafish is an excellent model for elucidating the genetic contribution to complications associated with anesthesia.

2. GENETIC MANIPULATION IN THE ZEBRAFISH

In recent years, there has been an explosion of technologies that make engineering the zebrafish genome both simple and reliable. These technologies are discussed below.

2.1. Reverse Genetics: Recapitulating Human Mutations in Zebrafish

Many inherited disorders are caused by nonsense, loss-of-function mutations in a specific gene. To generate a zebrafish model of this type of disorder, both knockdown and knockout approaches are used. To knockdown gene expression, morpholinos were developed in the late 1990s (Summerton & Weller, 1997). Morpholinos are antisense nucleotides primarily used to study genes expressed within the first 2–3 days postfertilization (dpf). At later stages, they are unreliable due to degradation and dilution. They are also not ideal for studying the p53 pathway and neurological development due to their nonspecific upregulation of p53 causing nonspecific neurologic damage. They are, however, very simple to design and to use as reviewed in Bedell, Westcot, and Ekker (2011) and Stainier et al. (2017).

Better suited to anesthesia studies are genome engineering approaches that make stably inherited changes to the genome. These methods create double-stranded breaks at a designated site in the genome (Fig. 1). The break is then repaired by endogenous machinery using nonhomologous end joining, which causes a small number of base pairs to be trimmed off or added to the ends of the break before those loose ends are fused back together (Bibikova, Golic, Golic, & Carroll, 2002). This generally creates a small insertion or deletion (an “indel”). These indels can cause loss-of-function, nonsense mutations resulting in the introduction of a stop codon and premature protein truncation. While it is possible to use zinc finger endonucleases (Doyon et al., 2008) or transcription activator-like effector nucleases (TALENs) (Huang et al., 2011), the most common approach currently is based on CRISPRs (clustered regularly interspaced short palindromic repeats)/Cas9. For this method, an endonuclease protein, Cas9, binds to a small guide RNA (sgRNA; Fig. 1D) to target a specific genomic sequence and create double-stranded breaks: reviewed in Hsu et al. (2014) and Varshney et al. (2016). The workflow for disrupting a gene is shown in Fig. 1.

Fig. 1.

Fig. 1

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Genetic manipulation of zebrafish by injecting 1-cell stage embryos. Fertilized eggs surrounded by an acellular chorion are collected and aligned against a glass slide taped to a petri plate (A). These are injected on a dissecting microscope equipped with a foot pedal-driven pressure injector and manipulator (B). For CRISPR/Cas9, small guide RNAs and Cas9 protein are mixed and loaded into an injection needle that is then mounted on the manipulator (C and D). Depending on the topic of study, different stable transgenic lines can be injected. As in Varshney et al. (2016), five guide RNAs are designed against a single exon of the target gene and coinjected with Cas9 protein. Subsequently, embryos are tested for genome editing by using a three-primer polymerase chain reaction to make fluorescent amplicons of the edited site. These are then run on a sequencing capillary to see either a series of insertions (in) and deletions (dels) in the embryo injected with the guide RNAs or a single wild-type band in the case of the embryos that are only injected with cas9 (E).

It is also possible to specifically engineer missense mutations that change a single, targeted base pair change resulting in a specific change in amino acid sequence. This was first accomplished in zebrafish using TALENs (Bedell et al., 2012) and then CRISPRs (Hoshijima, Jurynec, & Grunwald, 2016; Ran et al., 2013). The TALENs/CRISPRs are designed as described earlier to target a specific genetic locus. Then a single-stranded DNA oligo is designed with the mutation of interest flanked by homologous arms that are around 10 base pairs, with the entire oligo being around 20 base pairs. The ssDNA provides a template for the repair machinery of the cell to undergo homology directed repair rather than nonhomologous end joining (Bedell et al., 2012). Alternatively, a donor plasmid can be engineered with longer homologous arms (Hoshijima et al., 2016). In these ways, specific, disease-causing human mutations can be recapitulated in a zebrafish model.

2.2. Expression of Transgenes

In addition to loss-of-function nonsense and missense alleles, gain-of-function or overexpression experiments can also be used to study missense alleles that produce dominant gain-of-function phenotypes (Koenighofer et al., 2016). Once again, there are both transient and permanent methods. To transiently overexpress, mRNA of a gene is injected into the single cell of the zebrafish embryo. This mRNA is translated, depending on the design of the untranslated regions, for as long as 2 dpf (Hyatt & Ekker, 1999).

If experiments require longer duration of expression or for the expression to be in a specific tissue, transposons can be used to insert DNA into the genome. Transposons are proteins that interact with a pair of specific repeats to excise a piece of DNA from one place and insert that DNA randomly into another. Any sequence including transgenes with transcriptional activator and promoter sequences as well as the coding sequence of the protein to be expressed can be placed between the two transposase recognition sequences (Clark et al., 2011). This approach results in stable expression of the transgene.

2.3. Zebrafish Models of Disorders Linked to Anesthetic Complications

Many zebrafish models have already been made, among them models of narcolepsy (Prober, Rihel, Onah, Sung, & Schier, 2006; Yokogawa et al., 2007), glycine encephalopathy (Cui et al., 2005; Mongeon et al., 2008), epilepsy/autism (Grone et al., 2016; Grone, Qu, & Baraban, 2017; Hoffman et al., 2016; Scheldeman et al., 2017), and Parkinson’s (Soman et al., 2017). These and other models can be leveraged to better understand the genetic contributions to complications associated with anesthesia.

2.4. Forward Genetics: Unbiased Discovery of Genetic Variants That Confer Risk for Anesthesia

To identify new genetic variants that alter responses to anesthetics, there are several methods regularly employed by the zebrafish community to generate mutant lines in an unbiased way. These lines are then studied by investigators interested in specific phenotypes (Mullins, Hammerschmidt, Haffter, & Nusslein-Volhard, 1994). Traditionally, mutations are generated with chemicals, such as N-ethyl-N-nitrosourea (ENU), that create single base pair changes: reviewed in Knapik (2000). This method creates base pair changes in up to 1 in every 100 base pairs. Therefore, it can be difficult to assess which mutation is causative for the phenotype. Additionally, it can be difficult to find the mutation of interest because it is a single base pair change in an unknown gene. To address these issues, insertional mutagenesis has also been used including DNA viruses (Amsterdam et al., 1999) and transposons (Clark et al., 2011; Varshney et al., 2013). Similar forward genetic approaches in invertebrate models have defined new anesthetic targets (Morgan & Sedensky, 2013).

Several distinct large collections of mutant zebrafish lines are maintained at large facilities including the National Institutes of Health (Varshney et al., 2013), European Zebrafish Resource Center (EZRC) (Geisler, Borel, Ferg, Maier, & Strahle, 2016), Chinese Zebrafish Resource Center (https://zfin.org/ZDB-LAB-130226-1), and Zebrafish International Resource Center (Varga, 2016).

3. BEHAVIORS TO ASSESS SEDATION AND RECOVERY FROM ANESTHESIA

Zebrafish have emerged as the model organism of choice for high-throughput drug screening of behavioral phenotypes (Bruni, Lakhani, & Kokel, 2014; Kokel & Peterson, 2011; Rihel et al., 2010). A number of behaviors have been analyzed and reported using the zebrafish model including locomotion, feeding, cognition, memory, and stress (Kalueff et al., 2013; Orger & de Polavieja, 2017). However, in order to exploit the advantages of the zebrafish model and to achieve rapid and statistically robust readouts, it is essential to phenotype the behavioral patterns in high throughput. The Screening Center at the Karlsruhe Institute of Technology is a transdisciplinary center that specializes in developing automated and intelligent morphological and behavioral screening platforms (e.g., Figs. 3 and 4). Every platform is equipped with automated sampling handling systems, a high-content and high-throughput imaging component, an image analysis pipeline for real-time data analysis and finally a large-scale data storage system for data archival and retrieval. The center is connected with the EZRC that serves as a hub to obtain wild type, mutant and transgenic zebrafish and medaka. The Screening Center offers options to conduct morphological and behavioral screening for zebrafish and the medaka community. Here we report two behavioral phenotyping platforms that function in high throughput and can be used efficiently to understand and quantify effects of anesthetics.

Fig. 3.

Fig. 3

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PMR high-throughput pipeline. The upper panel describes the flow of the PMR experiment. (A) The imaging system with A, representing the power source;B, the shutter controller;C, the stage controller;D, the custom IR ring;and E, the microscope itself. (B) A snapshot of a single well with embryos that will be subjected to the light flash. (C) Representative results of the PMR response to 3 different compounds with 10 repeats for each compound showing typical PMR phenotypes: normal response on the top, the middle graph showing increased activity throughout the different PMR phases, and finally the bottom graph showing increased latency and high activity afterward.

Fig. 4.

Fig. 4

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High-throughput vibration analysis platform. (A) The screening platform showing five different vibration pulse sources mounted below five petri-dishes containing embryos. The robotic arm carries the high-speed camera that moves sequentially over the different source positions. (B) A typical C bend response after the application of the vibration pulse.

3.1. Photomotor Response (PMR)

The PMR is one of the earliest manifestations of behavior in zebrafish occurring between 30 and 42h hpf and does not involve any canonical visual organs (Kokel et al., 2013; Kokel & Peterson, 2011). Between 30 and 42 hpf, zebrafish embryos exhibit a low basal movement. The application of a strong and short pulse of light, however, evokes rapid movement of the embryos within the chorion after a few seconds of latency that lasts for about 5s. After this, the embryos enter a refractory phase and do not respond to a second pulse of light (Kokel & Peterson, 2011; Marcato et al., 2015). Fig. 2 shows this behavior schematically.

Fig. 2.

Fig. 2

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The photomotor response (PMR) of zebrafish embryos between 30 and 42hpf.

Interestingly, it has been found that this behavior is significantly altered in the presence of neuroactive compounds and anesthetics. Consequently, we have developed a high-throughput PMR platform that is completely automated from the sampling handling stage to the analysis stage. The entire platform functions sequentially and is described below and shown in Fig. 3 (Marcato et al., 2015).

Step 1: Sample handling

In the first step, zebrafish embryos are automatically sorted to remove coagulated and unfertilized eggs. After this, 10–12 embryos are automatically pipetted into the wells of a standard 96-well microtiter plate. Furthermore, the entire process works in the absence of ambient white light (thereby avoiding exposure to light before the experiment) as this system operates under infrared illumination. This step helps in avoiding human errors and facilitates the preparation of a large number of plates per day for experimentation.

Step 2: Data acquisition

After sample handling, the automatically prepared plate is transferred to a custom-built imaging system to conduct the actual PMR experiment. A Nikon Eclipse Ti-E inverted microscope equipped with a custom LED ring is programed so that each well containing embryos is successively in the focal plane of the microscope. The light flash is provided by a Nikon 100W Xenon bulb and a controllable shutter (Sutter Instruments, Novato, CA, USA). The video recording is done using a Nikon DS-Qi 1 monochrome camera with a 1280 × 1024 pixel resolution at 19 frames per second and with a CFI Plan Achromat UW 2 × objective to get a large field of view to cover the entire well. The entire acquisition and control software is custom developed.

Step 3: Data analysis and visualization

An entire analysis and visualization pipeline were developed to automatically analyze the data acquired from the PMR experiment. The analysis begins with egg detection using a circular Hough Transform and a weighting algorithm that avoids false detections. A motion index which is a numerical value reflecting the changes in consecutive frames of the PMR video is then calculated for each egg. The temporal change in the motion index is then reflective of the test compound on the PMR behavior.

Different anesthetics show different effects on the PMR including decreased activity on the application of a light pulse, increased time to respond (increased latency), and for some doses no response to the application of the light pulse. Taken together, this is a complex, noninvasive behavioral readout that can easily be used to assess anesthetic response and any changes in that response due to genomic changes.

3.2. Startle/Vibration Response

A startle response is a fast reaction to an applied vibrational stimulus or sound. At about 4 dpf zebrafish larvae respond to vibration in a characteristic way (Burgess & Granato, 2007). The maturation of the ear/swim bladder to Mauthner neuron escape circuit is believed to be responsible for this behavior: reviewed in Eaton, Lee, and Foreman (2001). With application of a vibration pulse the larvae respond by bending the head all the way to the tail forming a C shape. This is known as the C bend. The larvae then release themselves from this position and propel themselves forward and swim away in an escape response. This behavior is robust and consistent and like the PMR can be modulated by anesthetics and other neuroactive compounds. In the absence of a high-throughput vibration response system, we have developed one that automatically images the response and analyzes the data to provide kinematic parameters. We have used this system to study emergence time of zebrafish larvae subjected to different anesthetics and at different doses.

The system consists of several acoustic speakers that are calibrated to provide a vibration pulse of around 15g for 1ms. The response is then recorded using a high-speed camera at 1000 frames per second. The data are then automatically analyzed to extract activity, the bend angles and the distance traveled after the application of the pulse. The system is shown in Fig. 4.

4. ANESTHESIA IN ZEBRAFISH

Methods of delivering anesthetics vary depending on life-history stage. While gavage is used in adult zebrafish and other adult teleosts (Dang, Henderson, Garraway, & Zon, 2016), anesthetics are more often added directly to the bath (immersion) for larval zebrafish. Here, we focus on larval stages and immersion because these are the stages that are amenable to high-throughput behavioral studies.

4.1. Types of Anesthetics

There is a growing body of literature on the behavioral and physiologic effects of various anesthetics on teleosts (Collymore, Tolwani, Lieggi, & Rasmussen, 2014; Readman, Owen, Knowles, & Murrell, 2017; Sneddon, 2012). The most commonly used anesthetics, mechanisms of action, behavioral and physiologic effects on fish are described in many publications elsewhere and are beyond the scope of this review. Anesthetics evaluated in zebrafish range from nonpharmaceutical grade anesthetics such as MS222/benzocaine (Sneddon, 2012), Eugenol (Javahery, Nekoubin, & Moradlu, 2012), and carbon dioxide administered with ultrafine oxygen bubbles (Kugino, Tamaru, Hisatomi, & Sakaguchi, 2016) and pharmaceutical grade general anesthetics used in humans including isoflurane (Huang et al., 2010) and propofol (Warren, Baker, & Fishman, 2001). Isoflurane is also sometimes used in combination with MS222 in the zebrafish (Huang et al., 2010). The combination is preferable to MS222 alone because it does not depress cardiac function, fish can be sedated for longer periods of time, and recover faster. The same is true for propofol (Warren et al., 2001). The fact that propofol and isoflurane are effective in both zebrafish and humans raises the potential for the discovery of new anesthetics using high-throughput behavioral screening in zebrafish. Such a screening strategy was shown to be effective for identifying novel antipsychotics (Bruni et al., 2016).

4.2. Special Considerations When Making Propofol

Owing to its hydrophobic nature, propofol adheres to plastic and is not particularly water soluble. Therefore, to avoid uncertain concentrations due to loss from absorption, glass is used to pipette and store propofol. To break up micelles that form when propofol is added to aqueous solutions, it is essential to vortex and sonicate the solution (Fig. 5). A previous publication that used propofol in zebrafish larvae (Warren et al., 2001) used a 10-fold higher dose than what is known to anesthetize Xenopus tadpoles (Krasowski et al., 2001; Weiser, Kelz, & Eckenhoff, 2013) and humans (Purdon et al., 2013), likely due to these solubility issues. An alternate method is to use DMSO as a cosolvent or to use a formulation of propofol that is more readily soluble in water. When propofol solutions are made using the following protocol (Fig. 5), sedative (~1μM) and anesthetic (~10μM) doses of propofol for zebrafish are similar to what have been reported for tadpoles and humans (Fig. 6A and B).

Fig. 5.

Fig. 5

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Diluting propofol for anesthesia. To make propofol, we first use a Bunsen burner (A) to make a drawn out glass pipet that we calibrate to 1 μL (B). We add 1 μL propofol to 100mL system water in a glass bottle, vortex for 30s (C), and sonicate for 5 min (D).

Fig. 6.

Fig. 6

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Vibration response can be used to assess sedation and emergence parameters for anesthesia. (A) 10–20 five-day-old larvae are exposed to vibration for every 20s for 5min after addition of propofol and the proportion responding normalized to baseline is plotted vs time for different concentrations of propofol. (B) A dose–response curve is constructed from proportion responding at 5min. Proportions are fitted with a logistic regression in JMP13 (green line).

For our purposes, we:

  1. Make a drawn out glass pipet using a Bunsen burner (using hemostats; Fig. 5A)

  2. Break of the end of the glass pipet and dispose in glass waste

  3. Measure 1 μL water with green food coloring onto parafilm using a pipetman

  4. Draw up solution into newly fashioned glass pipet and mark the fill line (Fig. 5B)

  5. Wash pipet in water and then use to measure propofol

  6. Add 100mL system water to a 100mL glass bottle

  7. Pipet 1μL propofol into 100mL

  8. Vortex 30s (Fig. 5C)

  9. Sonicate 5min (Fig. 5D)

  10. Measure concentration on a spectrophotometer, measuring absorbance @270nm. The molar extinction coefficient is 1600/M.

4.3. HPLC to Determine Tissue Concentration of Anesthetic

Because the route of delivery of the propofol anesthetic is immersion, it is important to determine the concentration that has reached the brain. To do this we euthanize fish and then isolate brain tissue for HPLC using the following protocol:

  1. Prepare several 35-mm petri plates for surgery arena
    1. Coat with Sylgard (Corning)

    mix polymers 1:10

    pour ~3mm depth in each petri

    allow to harden overnight

    soak in distilled water before using for surgeries

  2. Expose 5–7-day-old larvae to anesthetic in baskets for ease of exposure to anesthetic and washing

  3. After anesthetic exposure, euthanize larvae by placing baskets on dry ice

  4. Place surgery arena in 10cm petri filled with wet ice

  5. Transfer basket from dry ice and use forceps to transfer euthanized larva by its tail to the surgery arena so that the head faces the inside of the plate and the notochord is visible

  6. Pin the larva to the plate using FST insect pins bent to a right angle with forceps

  7. Add a couple drops of system water over the larva
    1. System water is taken from recirculating aquaria housing adult fish and contains salt for an osmolarity between 400 and 500mOsm and a neutral pH ~7.4

  8. Remove the eyes using the tungsten needles (Electron Microscopy Sciences)

  9. Sever the head (usually do this right before the swim bladder)

  10. Remove the skin and ears from the head by running over the matter with the tungsten needles

  11. Using the tungsten needles drag the brain matter outside of the water using the surface tension to make a bubble on the needle with the brain matter inside

  12. Transfer to an Eppendorf and store on dry ice for HPLC as described in Gardner et al. (2016).

4.4. Propofol Sedation Curve

To construct dose–response curves (Fig. 6A and B), 10–20 five-day-old larvae are placed in 1.5mL system water in a 35mm petri dish lined with a sylgard ring to keep larvae in the field of view. This dish is placed on an aluminum ring attached to a minishaker by a titanium rod to deliver calibrated vibrational stimuli. Vibration-evoked escape responses (Burgess & Granato, 2007) require sensory/motor integration similar to the righting reflex used in rodents (Campagna, Miller, & Forman, 2003) and are therefore appropriate for assessing depth of sedation. In Fig. 6A, stimuli are delivered every 20s with responses coming to steady state by 4 min after addition of propofol. The dose–response curve (Fig. 6B) is constructed by taking proportion responding at 5 min normalized to averaged responsiveness of the larvae during a baseline test conducted before addition of propofol. This curve can also be obtained using PMR with zebrafish embryos at very early stages of development. These two assays, PMR (at 30hpf) and vibration response (at 4dpf), can help our understanding of how the response to anesthetics changes with age.

5. CONCLUSIONS

The zebrafish has become a powerful model for delineating the genes and neural circuits that contribute to a given behavior. Genome engineering approaches like CRISPR/Cas9 have made it straight forward to generate zebrafish models of human inherited disorders associated with anesthetic risk. In addition, collections of randomly mutated lines of zebrafish enable forward genetic screens for those genes as yet unknown that cause complications. The sensory-evoked and highly stereotyped zebrafish behaviors, such as the PMR and vibration response, can be studied in a high-throughput setting that allows for fast screening of both mutations and drugs. Furthermore, it has been shown that zebrafish larvae respond to propofol and other anesthetics in a similar pattern and concentration as other animals. Taken together, the zebrafish is poised to enhance our understanding of the mechanism of anesthesia and the role genetics plays in the response to anesthesia.

Key Points.

  1. Genetic tools available to knockout or overexpress genes include both transient and stably inheritable approaches. Specific base pair changes can be engineered within the genome. Additionally, forward genetic random mutation screening is readily available.

  2. Multiple complex, validated behavior assays are available to study drug response, including the photomotor and startle responses.

  3. The zebrafish is an idea model for high-throughput studies to discover new genes for anesthetic response or drugs that induce anesthesia.

  4. Owing to its hydrophobic nature, use of propofol with the zebrafish requires use of glass equipment and careful mixing into solution to maintain a consistent concentration.

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