PMCCPMCCPMCC

Search tips
Search criteria 

Advanced

 
Logo of nihpaAbout Author manuscriptsSubmit a manuscriptHHS Public Access; Author Manuscript; Accepted for publication in peer reviewed journal;
 
Dev Biol. Author manuscript; available in PMC 2014 May 8.
Published in final edited form as:
PMCID: PMC4013685
NIHMSID: NIHMS573066

Disruption of Eaat2b, a glutamate transporter, results in abnormal motor behaviors in developing zebrafish

Abstract

Analysis of zebrafish mutants that have defects in motor behavior can allow entrée into the hindbrain and spinal cord networks that control locomotion. Here, we report that zebrafish techno trousers (tnt) locomotor mutants harbor a mutation in slc1a2b, which encodes Eaat2b, a plasma membrane glutamate transporter. We used tnt mutants to explore the effects of impaired glutamate transporter activity on locomotor network function. Wild-type larvae perform robust swimming behavior in response to touch stimuli at two and four days after fertilization. In contrast, tnt mutant larvae demonstrate aberrant, exaggerated body bends beginning two days after fertilization and they are almost paralyzed four days after fertilization. We show that slc1a2b is expressed in glial cells in a dynamic fashion across development, which may explain the abnormal sequence of motor behaviors demonstrated by tnt mutants. We also show that tnt larvae demonstrate enhanced excitation of neurons, consistent with the predicted effects of excessive glutamate. These findings illustrate the dynamic regulation and importance of glutamate transporters during development. Since glutamate toxicity caused by EAAT2 dysfunction is thought to promote several different neurological disorders in humans, including epilepsy and neurodegenerative diseases, tnt mutants hold promise as a new tool to better understand these pathologies.

Keywords: Behavior, Zebrafish, Glutamate, Transporter, Hindbrain, Spinal cord

Introduction

The neural networks that coordinate motor behavior rely upon a variety of neurotransmitter systems, including glutamate. Glutamatergic neurotransmission is shaped and regulated by sodium-dependent plasma membrane transporters. These transporters control extracellular levels of glutamate, which can be toxic to neurons if present at high concentrations (Olney, 1969). In mammals, glutamate plasma membrane transporters are encoded by five genes: Slc1a1–5, which encode the proteins EAAT1–5. One of these transporters, EAAT2 (also known as GLT1), is expressed predominantly in glial cells (Lehre et al., 1995; Rothstein et al., 1994, 1995). EAAT2 plays a central role in glutamate uptake by providing ~90% of plasma membrane glutamate transport in adult tissue (Lehre et al., 1995). Although EAAT2 is expressed during central nervous system (CNS) development, the potential roles it plays during neural network formation have not been well examined.

The zebrafish, Danio rerio, provides an excellent system to study how Eaat2 transporters govern neural network development and function. Although they have not been fully described, the zebrafish orthologs are expressed in developing and adult animals (Gesemann et al., 2010; Rico et al., 2010; Rohrschneider et al., 2007). In addition, zebrafish embryos are readily amenable to a variety of imaging, pharmacological, genetic and behavioral approaches. Motor behaviors are performed in a characteristic sequence during development, which results in a robust escape response by 40 hours post-fertilization (hpf) (Buss and Drapeau, 2001; Kimmel et al., 1974; Saint-Amant and Drapeau, 1998). The escape response consists of a large amplitude body bend, known as a C-start, away from the stimulus, followed by smaller amplitude body undulations to swim away. Several physiology and lesion studies indicate that neural networks in the hindbrain, including the Mauthner cells and other reticulospinal neurons, can initiate the large amplitude body bend of the C-start, while spinal cord networks can generate the lower amplitude body undulations of swimming (Cox and Fetcho, 1996; Downes and Granato, 2006; Liu and Fetcho, 1999; Nissanov et al., 1990; Zottoli, 1977).

Here, we characterize the zebrafish techno trousers (tnt) behavioral mutant and reveal that the tnt mutation disrupts slc1a2b, which encodes Eaat2b. We then examine the expression of slc1a2b during development. Finally, we use tnt mutants to examine the in vivo effects of slc1a2b/eaat2b dysfunction on neuronal activity. Taken together, these data illustrate the central role that slc1a2b/eaat2b plays in regulating locomotor network function and identify the tnt mutant as a new in vivo system to examine the effects of impaired glutamate transport.

Material and methods

Fish maintenance and breeding

Zebrafish were raised and maintained as previously described (Mullins et al., 1994). Embryos were kept at 28.5 °C in E3 media and staged according to morphological criteria (Kimmel et al., 1995). All experiments were performed using k57 (tnt) or wild type embryos in a Tübingen (Tü) or tub longfin (TLF) genetic background. All animal protocols were approved by the University of Massachusetts or University of Colorado Institutional Animal Care and Use Committees (IACUC).

Behavioral analysis

A light tap to the head was applied with a blunted insect pin to elicit escape responses. Each response was recorded using a high-speed digital camera (Fastec Imaging, San Diego, CA), mounted to a 35 mm lens (Nikon, Melville, NY), collecting at a rate of 500 frames/s. The head-to-tail angle for each frame of the response was measured using automated software developed by our lab and plotted using Microsoft Excel. The duration of an escape response was calculated as beginning in the frame immediately before movement was first detected and lasting until the frame when movement was no longer observed.

Lesion experiments

Wild-type and mutant embryos were staged at ~46 hpf and placed in 1× Ringers solution (116 mM NaCl, 2.9 mM KCl, 1.8 mM CaCl2, and 5 mM HEPES, pH 7.2). Once equilibrated in Ringers for 20 min, the embryos were anesthetized using 0.04% MS-222 (Tricaine) and lesioned as previously described (Downes and Granato, 2006). The lesioned preparations were placed in a new dish of 1× Ringers and allowed to recover for 1 h prior to any measurements. Both intact embryos and isolated tails were kept in 1× Ringers for the duration of the experiment. Digital pictures were taken of a three-somite region dorsal to the yolk extension at 48 and 96 hpf. These were calibrated to acquire the actual length measurement (in mm).

Chromosomal mapping and cDNA cloning

Crosses between fish heterozygous for the tnt allele and WIK fish were used to generate a three-generation map cross. The mapping procedure and the WIK line were described previously (Knapik et al., 1996; Rauch et al., 2003). F2 tnt mutant embryos and wild-type siblings were collected, sorted based upon the 96 hpf phenotype and stored in methanol at −20 °C. Bulk segregant analysis was performed by the Zebrafish Mapping Facility at the University of Louisville, Kentucky. DNA was extracted from pools of 20 mutant and 20 wild-type embryos and analyzed using simple sequence length polymorphism (SSLP) markers distributed throughout the zebrafish genome. To determine the tnt linkage interval, several nearby markers on Chromosome 25, including z1462 and z28616, were used with DNA from 96 individual F2 mutant embryos from tnt mapping cross lines.

To clone slc1a2b, RNA was extracted from 96 hpf wild-type and mutant embryos and used for RT-PCR. The Accuscript RT-PCR system (Stratagene, La Jolla, CA) was used with gene specific primers to reverse transcribe and amplify slc1a2b cDNA in sections. The purified amplification products were sequenced at core sequencing facilities (GeneWiz, South Plainfield, NJ) and analyzed using MacVector 9.0. Independent RT-PCR reactions were performed multiple times to reduce the likelihood of amplification artifacts.

Morpholino analysis

Wild-type zebrafish embryos were pressure injected at the 1- to 4-cell stage with 4 ng of morpholino designed to block translation of slc1a2b or the standard control morpholino. This amount of morpholino was selected based upon dose–response experiments in which higher doses were found to generate morphological defects and/or lethality. The sequence of the translation blocking morpholino was 5′-TCGGCATCATCCACAACTGTCAGGC-3′. The control morpholino sequence was 5′-CCTCTTACCTCAGTTACAATTTATA-3′. The embryos were raised at 28 °C, and locomotive behavior was examined at 24, 48, and 96 hpf.

Whole-mount in situ hybridization and antibody staining

Antisense digoxigenin probes were generated against slc1a2b using a cDNA clone acquired from Open Biosystems (Huntsville, AL). Whole-mount, colorimetric in situ hybridization was performed using established protocols and examined using a compound microscope (Zeiss, Thornwood, NY) attached to a digital camera (Zeiss, Thornwood, NY).

slc1a2b mRNA and different cell type selective markers were examined in the same embryo using fluorescent in situ hybridization combined with antibody staining. Fluorescent whole-mount in situ hybridization and antibody staining were performed as previously described with few modifications (Downes and Granato, 2004). slc1a2b was visualized using Fast Red as the chromogenic substrate (Roche Ltd., Basel, Switzerland). GFAP:GFP and mnx1:GFP transgenic lines were used to examine glial cells and a population of motor and ventral interneurons, respectively (Bernardos and Raymond, 2006; Brand et al., 1996; Flanagan-Steet et al., 2005). GFP was detected using an anti-GFP antibody (1:20 dilution, Living colors peptide antibody, Clontech, Mountain View, CA). The 3A10 antibody was used to examine the Mauthner cells (Hatta, 1992). This antibody was obtained from the Developmental Studies Hybridoma Bank developed under the auspices of the NICHD and maintained by The University of Iowa, Department of Biological Sciences, Iowa City, IA 52242 (1:50 dilution). To visualize the primary antibodies, we used anti-rabbit Alexa 488 (1:500 dilution, Invitrogen, Inc., Carlsbad, CA) and anti-mouse Alexa 488 (1:500 dilution, Invitrogen, Inc., Carlsbad, CA).

Electrophysiology recordings

Wild-type sibling or tnt mnx1:GFP larvae were anesthetized in 0.02% tricaine and sacrificed. Prior to recordings, larvae were glued (3 M Vetbond, Revival Animal Health, Orange City, IA) to a sylgard chamber (Dow Corning, Midland, MI) and skinned while immersed in Ringer’s solution (in mM: 145 NaCl, 3 KCl, and 1.8 CaCl2, 10 HEPES) as described previously (Moreno and Ribera, 2010). Whole cell current-recordings were obtained from CaP motor neurons using patch electrodes (2.5- to 3.5-MΩ resistance) and an Axopatch-200B amplifier (Axon Instruments, Molecular Devices, Sunnyvale, CA). Electrodes were made using a P-97 microelectrode puller (Sutter Instruments, Novato, CA), and filled with pipet solution containing (in mM: 105 K- gluconate, 16 KCl, 2 MgCl2, 10 HEPES, 10 EGTA and 4 NaATP; pH 7.2). During current-clamp recordings larvae were perfused with an external solution containing (in mM: 134 NaCl, 2.9 KCl, 2.1 CaCl2, 1.2 MgCl2, 10 HEPES and 10 Glucose; pH 7.8) and paralyzed with 0.4–0.8 μM α-Bgtx (Tocris, Ellisville, MO). Muscle from two hemisegments was removed by aspiration and meninges were carefully pierced to expose CaP motor neurons for recordings at 72 and 96 hpf. Only one or two hemisegments, at positions 3–6 in the rostral spinal cord were analyzed for each larva. CaP motor neurons were easily identified by their GFP expression driven by the mnx1 promoter and more dorsal soma position relative to CaP-like secondaries. CaP motor neurons were filled with a red fluorescent dye (100 μM Alexa Fluor 594; Invitrogen, Eugene, OR) during recordings to confirm their identity. Bursts were elicited by 30 ms supraspinal electrical stimulation at intensities ranging between 50 and 700 μA delivered through an electrode filled with extracellular solution (STG 1002, Multichannel Systems, ALA Scientific Instruments, Farmingdale, NY). Stimulation electrodes were fabricated using a Sutter P-97 microelectrode puller and broken to a diameter of ~10–20 μm by gentle brushing against the bottom of the recording chamber. In some experiments, the EAAT2 selective glutamate uptake blocker dihydrokainic acid (DHKA; Tocris, Ellisville, MO) was added at a concentration of 160 μM. Clampex 9.2 (Molecular Devices, Sunnyvale, CA) was used for data acquisition and analysis was performed with Clampfit 9.2 (Molecular Devices) and Axograph X (AxoGraph Scientific, Sydney, Australia). Burst durations were measured as the time from the end of stimulation artifact (erased from recordings shown) to the end of action potential firing when the membrane potential recovered to within 10 mV of its pre-burst value.

Results

tnt mutants demonstrate a progressive motility defect

A single allele of tnt, tk57, was isolated from a previously performed mutagenesis screen (Granato et al., 1996). Consistent with the initial description of the phenotype, no abnormalities were detected in early behavior, but beginning around 42 hpf tnt homozygous mutants display abnormal swimming. In response to touch stimuli, wild-type larvae perform the characteristic large-amplitude body bend of the C-start, followed by smaller amplitude body undulations to swim away (Fig. 1A; Movie 1; Fig. 4A). Similar to wild-type larvae, tnt mutants perform the initial large amplitude body bend however the smaller amplitude undulations are interrupted by exaggerated body-bends in which the nose touches the tail (Fig. 1B; Movie 2; Fig. 4B). This abnormal behavior sometimes appears to be struggling behavior, in which body undulations move in the direction opposite that from the rostral to caudal undulations of swimming behavior (Movie 2) (Liao and Fetcho, 2008). The duration of mutant escape responses is also longer than wild-type escape responses (Fig. 4F). By 96 hpf, wild-type larvae perform more rapid escape behavior but tnt larvae are nearly paralyzed. tnt mutants do twitch in response to touch, indicating that they can sense touch stimuli but cannot swim. Morphologically, mutants are shorter along the rostral–caudal axis compared to wild-type larvae (Fig. 2). tnt mutants fail to inflate the swim bladder and die around 6 days post-fertilization (dpf).

Fig. 1
tnt larvae exhibit an abnormal escape response at 48 hpf. Selected frames from high-speed video recordings are shown with times indicated in each frame in milliseconds. (A) A wild-type larva exhibits a normal C-bend (A2 — asterisk) in response ...
Fig. 2
tnt larvae demonstrate paralysis and body shortening at 96 hpf, which can occur in the absence of brain input. Lateral views are shown of wild-type and tnt larvae and lesioned preparations. (A,B) Wild-type and tnt larvae are of similar length along the ...
Fig. 4
slc1a2b knockdown phenocopies tnt mutants at 48 and 96 hpf. Kinematic traces are shown, with 0° indicating a straight body and positive and negative angles representing body bends in opposite directions. Time is shown in seconds. Five representative ...

To investigate whether the body shortening aspect of the tnt phenotype is dependent upon brain input to the spinal cord or can be mediated by only spinal cord networks, we performed gross lesion analysis. We transected 46 hpf wild-type and tnt mutant larvae just caudal to the otic placode to remove hindbrain input. Our previous work has shown that trunks isolated from transected embryos can be maintained for multiple days and demonstrate robust locomotive behavior (Downes and Granato, 2006). Mutant and wild-type preparations were indistinguishable from each other at 46 hpf and responded to touch with similar response durations (mutant: 281±45 ms, n=9; wild type: 359±145 ms, n=10; p=0.62, two-tailed t-test) (Movie 3, 4). These preparations were maintained and examined at 96 hpf. tnt preparations were paralyzed and significantly shorter along the rostral-caudal axis compared to wild-type controls (p=0.03) (Fig. 2E and E–H). These results suggest that while earlier aspects of the tnt phenotype may require brain input, the body shortening component of the phenotype becomes intrinsic to the spinal cord by 96 hpf.

The tnt gene encodes Eaat2b, a glutamate transporter

To determine the molecular identity of tnt, we first mapped the mutation using a three-generation map cross panel. Genomic DNA, from pooled and individual wild-type and homozygous mutant larvae, was screened with a panel of simple sequence length polymorphism markers. We found that tnt maps to a telomeric region of chromosome 25, close to markers z28616 and z14522 (0 out of 93 embryos for each marker) (Fig. 3A). Database analysis of candidate genes within this region revealed slc1a2b, which encodes Eaat2b, a plasma membrane glutamate transporter. Glutamate transporters have been identified in a variety of eukaryotic and prokaryotic species on the basis of four recognized sequence motifs (Slotboom et al., 1999). In mammalian systems, EAAT2 is predominantly expressed in glial cells within the central nervous system, and it plays a crucial role in removing glutamate from synapses (Lehre et al., 1995; Rothstein et al., 1994; Rothstein et al., 1995).

Fig. 3
The tnt gene encodes a glutamate transporter, Eaat2b. (A) The tnt locus was mapped to a telomeric portion of chromosome 25 that contains the slc1a2b gene, which encodes Eaat2b. Meiotic markers are shown at left and some of the neighboring genes are shown ...

Given the importance of EAAT2 for regulating glutamate levels in synaptic regions, we investigated if slc1a2b contained the tnt mutation. We cloned and sequenced slc1a2b transcripts from wild-type and homozygous mutant larvae. A single polymorphism was identified that substitutes a Val residue for Ala393 (Fig. 3B). Ala393 is contained within one of the four recognized glutamate transporter sequence motifs and it is precisely conserved across species from Escherichia coli to mammals (Fig. 3C). Ala393 also lies within a transmembrane domain that is essential for transporter activity (Fig. 3D) (Vandenberg et al., 1995; Yernool et al., 2004). In fact, mutation of the homologous residue from rat (Ala407) in cysteine scanning mutagenesis experiments led to a roughly 65% decrease in substrate transport (Zarbiv et al., 1998), consistent with the tnt mutation being a loss-of-function allele that diminishes or abolishes Eaat2b function.

To confirm the molecular identity of tnt, we injected antisense morpholino oligonucleotides (MO), designed to block translation of Eaat2b, into wild-type embryos at the 1–4 cell stage. No abnormalities were detected in early behavior, however at 48 hpf most slc1a2b morphant larvae (69 of 82 injected embryos, 84%) demonstrated a marked increase in the number of C-like body bends and prolonged swimming episodes, similar to tnt mutants (Fig. 4). In contrast, embryos injected with the control MO demonstrated behavior similar to wild-type embryos (n=73) (Fig. 4C,E, and F). Similar to tnt mutants, slc1a2b morphant larvae also demonstrated impaired swimming ability and statistically significant body shortening at 96 hpf (Fig. 4G). To further confirm the molecular identity of tnt we also attempted rescue analysis by mRNA injection at the 1–4 cell stage, but the late appearance of the tnt phenotypes likely hindered these experiments (data not shown). However, the mapping data, nature of the amino acid substitution, and MO phenocopy all argue that the tnt mutation disrupts slc1a2b/Eaat2b function.

slc1a2b exhibits dynamic expression in glial cells during development

Given the importance of hindbrain and spinal cord networks for the escape response and the abnormal escape behavior of tnt mutants, we expected to detect slc1a2b mRNA expression in these regions of the nervous system by 48 hpf. slc1a2b expression was examined using whole-mount in situ hybridization. Strikingly, we detected expression as early as 19 hpf in a portion of the hindbrain, rhombomere 4 (Fig. 5A and B). Rhombomere 4 contains the Mauthner cells, paired reticulospinal neurons that can initiate the C-start escape response (Eaton et al., 1977; Gahtan et al., 2002; Liu and Fetcho, 1999; Takahashi et al., 2002). Between 24 and 96 hpf, slc1a2b expression becomes more widespread and expands to many brain regions and the spinal cord (Fig. 5C–H). The pattern of expression, combined with results obtained from lesion analysis, suggests that the CNS domains that require slc1a2b function change and expand during development.

Fig. 5
slc1a2b exhibits dynamic expression in the hindbrain and spinal cord across development. Lateral (A,C,E,G) and dorsal views (B,D,F,H) are shown, with the brackets indicating the regions shown at higher magnification in the insets. For orientation in dorsal ...

In mammalian systems, Slc1a2 is expressed mainly in glial cells (Furuta et al., 1997; Lehre et al., 1995; Rothstein et al., 1994). To identify the cell types that express slc1a2b in larval zebrafish, we examined the mRNA distribution in comparison to markers for glial cells and two types of neurons that are known to mediate escape behavior: the Mauthner cells and motor neurons. We used the transgenic line Tg(gfap:GFP)mi2001/+ to examine slc1a2b expression in glial cells, the 3A10 antibody to examine expression in Mauthner cells, and the transgenic line Tg(mnx1:GFP)ml2/+ to examine expression in motor neurons (Bernardos and Raymond, 2006; Brand et al., 1996; Flanagan-Steet et al., 2005). At 48 hpf, slc1a2b is expressed throughout the hindbrain but not in Mauthner cells (Fig. 6A–F). Within the spinal cord, little colocalization is observed between slc1a2b mRNA and GFP+ cells in Tg(mnx1:GFP) embryos (Fig. 6G–I). In contrast, some colocalization was observed between slc1a2b mRNA and GFP+ cells in Tg(gfap:GFP) embryos, indicating that a subset of glial cells express slc1a2b (Fig. 6J–L). This analysis was also performed using 96 hpf larvae and very similar results were obtained (data not shown). Although we cannot eliminate the possibility of some neuronal expression, this analysis suggests that slc1a2b is expressed in at least some glial cells.

Fig. 6
slc1a2b mRNA is expressed in glial cells at 48 hpf. (A) The 3A10 antibody labels the Mauthner cell in the hindbrain. The scale bar=50 μm. (B) slc1a2b mRNA is expressed in the hindbrain in cells around the Mauthner cell. (C) The merged images show ...

Motor neurons in tnt mutants demonstrate abnormal bursting activity

The behavioral defects produced by the tnt mutation suggest abnormal neuronal activity. Reticulospinal neurons in the hindbrain, such as the Mauthner cells, can activate motor neurons in the spinal cord via glutamatergic synapses (Korn and Faber, 2005). Therefore we used supraspinal electrical stimulation combined with electrophysiological recording from spinal cord caudal primary (CaP) motor neurons to evaluate motor neuron activity and responses (Fetcho and Svoboda, 1993; McLean and Fetcho, 2009). Importantly, we tested for effects using intact preparations rather than dissociated cells to allow any effects of the tnt mutation on synaptic function to persist. In 72–96 hpf tnt larvae, CaP neurons have prolonged bursts compared to those in wild-type (mutant: 4370±560 ms; wild-type: 1460±210 ms; p=0.0001, two-tailed t-test) (Fig. 7A and B). The increase in burst duration in tnt mutants is consistent with an increase in the levels of extracellular glutamate produced by reduced or abolished glutamate transporter function. To determine whether direct blockade of Eaat2 transporters would result in similar effects, we used the selective blocker DHKA (160 μM; (Arriza et al., 1994; Wang et al., 1998). In 72–96 hpf wild type embryos, acute blockade of Eaat2 by DKHA results in an increase in burst duration (Fig. 7C and D), similar to the effect of the tnt mutation. Burst duration before and after DHKA shows significant differences (mean burst durations: before DHKA, 775±278 ms; after DHKA, 1274±309 ms; p=0.004, two-tailed pair t-test). Addition of CNQX (6-cyano-7-nitroquinoxaline-2,3-dione, 50 μM; MP Biomedicals, Solon, OH) and DL-2-amino-5-phosphonovaleric acid (AP-5, 100 μM; Tocris, Ellisville, MO) eliminated stimulated bursts, demonstrating that they resulted from activation of glutamate receptors (Buss and Drapeau, 2001) (data not shown).

Fig. 7
Motor neurons in tnt mutants exhibit increased burst duration. (A,B) Supraspinal electrical stimulation was used to elicit bursts in CaP motor neurons to evaluate synaptic input in 72–96 hpf larvae. Burst duration was increased in tnt CaP neurons ...

Discussion

In this study, we used the tnt mutant to examine the roles of slc1a2b/Eaat2b in regulating locomotor network function in developing zebrafish. The molecular nature of the tnt mutation indicates a loss or reduction of Eaat2b function. In vivo electrophysiology shows that slc1a2b/Eaat2b dysfunction results in enhanced excitation of motor neurons, which is consistent with the elevated amounts of extracellular glutamate predicted to result from impaired glutamate transporter function.

The lesion experiments and expression analysis suggest a model for how reduced or abolished Eaat2b function results in the abnormal sequence of behaviors demonstrated by tnt mutants. Early expression of slc1a2b is predominantly in rhombomere 4 of the hindbrain. This is the precise location of the Mauthner cells, which receive glutamatergic input to initiate the large amplitude body bend of a C-start (Diamond and Huxley, 1968; Korn and Faber, 2005). Moreover, Mauthner cells become functional around 40–48 hpf (Eaton et al., 1977), at which time tnt mutants first display abnormal behavior. Mauthner cells typically fire only once during an escape response and impaired Eaat2b function may result in abnormal excitation of Mauthner cells, resulting in the exaggerated body bends and prolonged swimming bouts that characterize tnt mutants.

Although slc1a2b mRNA is present in the spinal cord at 48 hpf, lesioning experiments suggest that it is not essential in this region of the CNS region until later in development. At 96 hpf, the tnt mutation no longer results in exaggerated body bends, but instead produces near paralysis and body shortening independent of hindbrain input. Exaggerated body bends, that are driven by abnormal hindbrain activity at 48 hpf, may not be observed at 96 hpf because hindbrain activity is now funneled through a severely disrupted spinal cord. At this time point, Eaat2b may be essential to control local spinal cord circuits. Diminished transporter function would be predicted to result in increased levels of extracellular glutamate. Such elevations in glutamate might lead to the increased motor neuron bursting, abnormal muscle contractions, and body shortening that we observe in tnt mutants.

Whereas excess glutamate would lead to increased excitation, another way to achieve excessive muscle contraction would be to reduce inhibition. Previous studies have shown that bandoneon mutants harbor a loss-of-function mutation in an inhibitory glycine receptor expressed within the hindbrain and spinal cord. Consistent with this idea, bandoneon mutants demonstrate a paralysis and body shortening phenotype very similar to tnt mutants at 96 hpf (Hirata et al., 2005).

Glutamate transporters are essential for neural development

Our data suggest that the spinal cord does not have an essential requirement for Eaat2b until after 48 hpf. However, glutamatergic neurotransmission occurs in the spinal cord prior to this time (Buss and Drapeau, 2001; Higashijima et al., 2004; Pietri et al., 2009). It is possible that glutamate re-uptake is not an essential mechanism to control synaptic glutamate levels at early stages of development as has been suggested in mammalian systems (Thomas et al., 2011). Alternatively, zebrafish contain several other glutamate transporters that may initially compensate for diminished Eaat2b function (Gesemann et al., 2010).

Studies in mouse provide evidence that glutamate transporters can partially compensate for each other. Targeted disruption of Slc1a2 did not produce any obvious defects at birth but about half of the animals died before 6 weeks of age due to severe spontaneous epileptic seizures (Tanaka et al., 1997). Although Eaat2 is responsible for the vast majority of extracellular glutamate reuptake in the adult brain, another glutamate transporter predominantly expressed in glial cells, Slc1a3/Eaat1 (also known as GLAST), is widely expressed during development (Matsugami et al., 2006; Shibata et al., 1996). Targeted disruption of Slc1a3/Eaat1 resulted in relatively mild defects. The Eaat1 deficient mice exhibited normal rates of survival, subtle motor defects, and increased susceptibility to cerebellar injury (Watase et al., 1998). In contrast to the results obtained when only one of the transporters was inactivated, severe defects in cell proliferation, migration and survival were observed throughout the brain when both Slc1a3/EAAT1 and Slc1a2/EAAT2 were deleted, demonstrating that these two transporters can partially compensate for each other during development (Matsugami et al., 2006).

tnt mutants as a model for disease

Disruption of SLC1A2/EAAT2 function has been implicated in several human neurological disorders, including epilepsy and amyotrophic lateral sclerosis (ALS). Alterations in SLC1A2 expression and mRNA splicing are observed in human patients with epilepsy, consistent with the epileptic seizures observed in Slc1a2 knockout mice (Rothstein et al., 1995; Tanaka et al., 1997). In the majority of ALS patients, substantial reductions in EAAT2 protein and concomitant increased levels of glutamate in cerebrospinal fluid have been found. No significant loss of glial cells was observed in these patients, indicating that decreased EAAT2 expression is not due to glial cell death (Rothstein et al., 1995). These data have led to the idea that reduced EAAT2 function results in elevated, toxic amounts of glutamate, which promote the motor neuron degeneration that is a central feature of ALS (reviewed in: (Fray et al., 1998; Maragakis and Rothstein, 2004; Sheldon and Robinson, 2007; Tanaka et al., 1997).

Zebrafish tnt mutants may be a new tool to examine the effects of excess glutamate generated by impaired glutamate transporter function. tnt mutants provide an accessible, in vivo system to delineate the cell type specific responses to glutamate toxicity. In addition, the tnt behavioral phenotype is robust, quantifiable and apparent very early in development compared to what has been detected in Slc1a2 deficient mice. Since zebrafish embryos are small, aquatic, available in large numbers, and develop outside of the mother, tnt mutants may be useful in chemical screens to identify small molecules that can ameliorate the effects of elevated glutamate (Zon and Peterson, 2010). This approach could be coupled with a variety of genetic, imaging, and physiological tools available in zebrafish. Taken together, these studies may provide a platform to identify drugs and genes to treat human diseases promoted by diminished EAAT2 function.

Acknowledgments

We thank the Karlstrom and Jensen labs for helpful discussion and Alexandra Nowlan for technical assistance. We also thank Saunders Whittlesey for kinematic analysis software development and Ronald Gregg and Greg Willer for help with mapping (supported by R01RR-020357). This work was supported by the National Institute of Neurological Disorders and Stroke grants F32NS-059120 to R.L.M., R01NS-025217, R01NS-038937, P30NS-048154 to A.B.R., and K01NS-057409 to G.B.D.

Footnotes

Supplementary materials related to this article can be found online at doi:10.1016/j.ydbio.2011.11.001.

References

  • Arriza JL, Fairman WA, Wadiche JI, Murdoch GH, Kavanaugh MP, Amara SG. Functional comparisons of three glutamate transporter subtypes cloned from human motor cortex. J Neurosci. 1994;14:5559–5569. [PubMed]
  • Bernardos RL, Raymond PA. GFAP transgenic zebrafish. Gene Expr Patterns. 2006;6:1007–1013. [PubMed]
  • Brand M, Heisenberg CP, Jiang YJ, Beuchle D, Lun K, Furutani-Seiki M, Granato M, Haffter P, Hammerschmidt M, Kane DA, Kelsh RN, Mullins MC, Odenthal J, van Eeden FJ, Nusslein-Volhard C. Mutations in zebrafish genes affecting the formation of the boundary between midbrain and hindbrain. Development. 1996;123:179–190. [PubMed]
  • Buss RR, Drapeau P. Synaptic drive to motoneurons during fictive swimming in the developing zebrafish. J Neurophysiol. 2001;86:197–210. [PubMed]
  • Cox KJ, Fetcho JR. Labeling blastomeres with a calcium indicator: a non-invasive method of visualizing neuronal activity in zebrafish. J Neurosci Methods. 1996;68:185–191. [PubMed]
  • Diamond J, Huxley AF. The activation and distribution of GABA and l-glutamate receptors on goldfish Mauthner neurones: an analysis of dendritic remote inhibition. J Physiol. 1968;194:669–723. [PubMed]
  • Downes GB, Granato M. Acetylcholinesterase function is dispensable for sensory neurite growth but is critical for neuromuscular synapse stability. Dev Biol. 2004;270:232–245. [PubMed]
  • Downes GB, Granato M. Supraspinal input is dispensable to generate glycine-mediated locomotive behaviors in the zebrafish embryo. J Neurobiol. 2006;66:437–451. [PubMed]
  • Eaton RC, Farley RD, Kimmel CB, Schabtach E. Functional development in the Mauthner cell system of embryos and larvae of the zebra fish. J Neurobiol. 1977;8:151–172. [PubMed]
  • Fetcho JR, Svoboda KR. Fictive swimming elicited by electrical stimulation of the midbrain in goldfish. J Neurophysiol. 1993;70:765–780. [PubMed]
  • Flanagan-Steet H, Fox MA, Meyer D, Sanes JR. Neuromuscular synapses can form in vivo by incorporation of initially aneural postsynaptic specializations. Development. 2005;132:4471–4481. [PubMed]
  • Fray AE, Ince PG, Banner SJ, Milton ID, Usher PA, Cookson MR, Shaw PJ. The expression of the glial glutamate transporter protein EAAT2 in motor neuron disease: an immunohistochemical study. Eur J Neurosci. 1998;10:2481–2489. [PubMed]
  • Furuta A, Rothstein JD, Martin LJ. Glutamate transporter protein subtypes are expressed differentially during rat CNS development. J Neurosci. 1997;17:8363–8375. [PubMed]
  • Gahtan E, Sankrithi N, Campos JB, O’Malley DM. Evidence for a widespread brain stem escape network in larval zebrafish. J Neurophysiol. 2002;87:608–614. [PubMed]
  • Gesemann M, Lesslauer A, Maurer CM, Schonthaler HB, Neuhauss SC. Phylogenetic analysis of the vertebrate excitatory/neutral amino acid transporter (SLC1/EAAT) family reveals lineage specific subfamilies. BMC Evol Biol. 2010;10:117. [PMC free article] [PubMed]
  • Granato M, van Eeden FJ, Schach U, Trowe T, Brand M, Furutani-Seiki M, Haffter P, Hammerschmidt M, Heisenberg CP, Jiang YJ, Kane DA, Kelsh RN, Mullins MC, Odenthal J, Nusslein-Volhard C. Genes controlling and mediating locomotion behavior of the zebrafish embryo and larva. Development. 1996;123:399–413. [PubMed]
  • Hatta K. Role of the floor plate in axonal patterning in the zebrafish CNS. Neuron. 1992;9:629–642. [PubMed]
  • Higashijima S, Mandel G, Fetcho JR. Distribution of prospective glutamatergic, glycinergic, and GABAergic neurons in embryonic and larval zebrafish. J Comp Neurol. 2004;480:1–18. [PubMed]
  • Hirata H, Saint-Amant L, Downes GB, Cui WW, Zhou W, Granato M, Kuwada JY. Zebrafish bandoneon mutants display behavioral defects due to a mutation in the glycine receptor beta-subunit. Proc Natl Acad Sci U S A. 2005;102:8345–8350. [PubMed]
  • Kimmel CB, Patterson J, Kimmel RO. The development and behavioral characteristics of the startle response in the zebra fish. Dev Psychobiol. 1974;7:47–60. [PubMed]
  • Kimmel CB, Ballard WW, Kimmel SR, Ullmann B, Schilling TF. Stages of embryonic development of the zebrafish. Dev Dyn. 1995;203:253–310. [PubMed]
  • Knapik EW, Goodman A, Atkinson OS, Roberts CT, Shiozawa M, Sim CU, Weksler-Zangen S, Trolliet MR, Futrell C, Innes BA, Koike G, McLaughlin MG, Pierre L, Simon JS, Vilallonga E, Roy M, Chiang PW, Fishman MC, Driever W, Jacob HJ. A reference cross DNA panel for zebrafish (Danio rerio) anchored with simple sequence length polymorphisms. Development. 1996;123:451–460. [PubMed]
  • Korn H, Faber DS. The Mauthner cell half a century later: a neurobiological model for decision-making? Neuron. 2005;47:13–28. [PubMed]
  • Lehre KP, Levy LM, Ottersen OP, Storm-Mathisen J, Danbolt NC. Differential expression of two glial glutamate transporters in the rat brain: quantitative and immunocytochemical observations. J Neurosci. 1995;15:1835–1853. [PubMed]
  • Liao JC, Fetcho JR. Shared versus specialized glycinergic spinal interneurons in axial motor circuits of larval zebrafish. J Neurosci. 2008;28:12982–12992. [PMC free article] [PubMed]
  • Liu KS, Fetcho JR. Laser ablations reveal functional relationships of segmental hindbrain neurons in zebrafish. Neuron. 1999;23:325–335. [PubMed]
  • Maragakis NJ, Rothstein JD. Glutamate transporters: animal models to neurologic disease. Neurobiol Dis. 2004;15:461–473. [PubMed]
  • Matsugami TR, Tanemura K, Mieda M, Nakatomi R, Yamada K, Kondo T, Ogawa M, Obata K, Watanabe M, Hashikawa T, Tanaka K. From the cover: indispensability of the glutamate transporters GLAST and GLT1 to brain development. Proc Natl Acad Sci U S A. 2006;103:12161–12166. [PubMed]
  • McLean DL, Fetcho JR. Spinal interneurons differentiate sequentially from those driving the fastest swimming movements in larval zebrafish to those driving the slowest ones. J Neurosci. 2009;29:13566–13577. [PMC free article] [PubMed]
  • Moreno RL, Ribera AB. Developmental regulation of subtype-specific motor neuron excitability. Ann N Y Acad Sci. 2010;1198:201–207. [PubMed]
  • Mullins MC, Hammerschmidt M, Haffter P, Nusslein-Volhard C. Large-scale mutagenesis in the zebrafish: in search of genes controlling development in a vertebrate. Curr Biol. 1994;4:189–202. [PubMed]
  • Nissanov J, Eaton RC, DiDomenico R. The motor output of the Mauthner cell, a reticulospinal command neuron. Brain Res. 1990;517:88–98. [PubMed]
  • Olney JW. Brain lesions, obesity, and other disturbances in mice treated with monosodium glutamate. Science. 1969;164:719–721. [PubMed]
  • Pietri T, Manalo E, Ryan J, Saint-Amant L, Washbourne P. Glutamate drives the touch response through a rostral loop in the spinal cord of zebrafish embryos. Dev Neurobiol. 2009;69 [PMC free article] [PubMed]
  • Rauch GJ, Lyons DA, Middendorf I, Friedlander B, Arana N, Reyes T, Talbot WS. Submission and Curation of Gene Expression Data. ZFIN Direct Data Submission 2003
  • Rico EP, de Oliveira DL, Rosemberg DB, Mussulini BH, Bonan CD, Dias RD, Wofchuk S, Souza DO, Bogo MR. Expression and functional analysis of Na(+)-dependent glutamate transporters from zebrafish brain. Brain Res Bull. 2010;81:517–523. [PubMed]
  • Rohrschneider MR, Elsen GE, Prince VE. Zebrafish Hoxb1a regulates multiple downstream genes including prickle1b. Dev Biol. 2007;309:358–372. [PubMed]
  • Rothstein JD, Martin L, Levey AI, Dykes-Hoberg M, Jin L, Wu D, Nash N, Kuncl RW. Localization of neuronal and glial glutamate transporters. Neuron. 1994;13:713–725. [PubMed]
  • Rothstein JD, Van Kammen M, Levey AI, Martin LJ, Kuncl RW. Selective loss of glial glutamate transporter GLT-1 in amyotrophic lateral sclerosis. Ann Neurol. 1995;38:73–84. [PubMed]
  • Saint-Amant L, Drapeau P. Time course of the development of motor behaviors in the zebrafish embryo. J Neurobiol. 1998;37:622–632. [PubMed]
  • Sheldon AL, Robinson MB. The role of glutamate transporters in neurodegenerative diseases and potential opportunities for intervention. Neurochem Int. 2007;51:333–355. [PMC free article] [PubMed]
  • Shibata T, Watanabe M, Tanaka K, Wada K, Inoue Y. Dynamic changes in expression of glutamate transporter mRNAs in developing brain. Neuroreport. 1996;7:705–709. [PubMed]
  • Slotboom DJ, Konings WN, Lolkema JS. Structural features of the glutamate transporter family. Microbiol Mol Biol Rev. 1999;63:293–307. [PMC free article] [PubMed]
  • Takahashi M, Narushima M, Oda Y. In vivo imaging of functional inhibitory networks on the mauthner cell of larval zebrafish. J Neurosci. 2002;22:3929–3938. [PubMed]
  • Tanaka K, Watase K, Manabe T, Yamada K, Watanabe M, Takahashi K, Iwama H, Nishikawa T, Ichihara N, Kikuchi T, Okuyama S, Kawashima N, Hori S, Takimoto M, Wada K. Epilepsy and exacerbation of brain injury in mice lacking the glutamate transporter GLT-1. Science. 1997;276:1699–1702. [PubMed]
  • Thomas CG, Tian H, Diamond JS. The relative roles of diffusion and uptake in clearing synaptically released glutamate change during early postnatal development. J Neurosci. 2011;31:4743–4754. [PMC free article] [PubMed]
  • Vandenberg RJ, Arriza JL, Amara SG, Kavanaugh MP. Constitutive ion fluxes and substrate binding domains of human glutamate transporters. J Biol Chem. 1995;270:17668–17671. [PubMed]
  • Wang GJ, Chung HJ, Schnuer J, Lea E, Robinson MB, Potthoff WK, Aizenman E, Rosenberg PA. Dihydrokainate-sensitive neuronal glutamate transport is required for protection of rat cortical neurons in culture against synaptically released glutamate. Eur J Neurosci. 1998;10:2523–2531. [PubMed]
  • Watase K, Hashimoto K, Kano M, Yamada K, Watanabe M, Inoue Y, Okuyama S, Sakagawa T, Ogawa S, Kawashima N, Hori S, Takimoto M, Wada K, Tanaka K. Motor discoordination and increased susceptibility to cerebellar injury in GLAST mutant mice. Eur J Neurosci. 1998;10:976–988. [PubMed]
  • Yernool D, Boudker O, Jin Y, Gouaux E. Structure of a glutamate transporter homologue from Pyrococcus horikoshii. Nature. 2004;431:811–818. [PubMed]
  • Zarbiv R, Grunewald M, Kavanaugh MP, Kanner BI. Cysteine scanning of the surroundings of an alkali-ion binding site of the glutamate transporter GLT-1 reveals a conformationally sensitive residue. J Biol Chem. 1998;273:14231–14237. [PubMed]
  • Zon LI, Peterson R. The new age of chemical screening in zebrafish. Zebrafish. 2010;7:1. [PubMed]
  • Zottoli SJ. Correlation of the startle reflex and Mauthner cell auditory responses in unrestrained goldfish. J Exp Biol. 1977;66:243–254. [PubMed]