A particular challenge in the design and execution of behavioral screens is the variability of normal behavior. In order to minimize environmental contributions to behavioral variability, a major effort must be devoted to controlling the raising and testing conditions. Before carrying out a genetic screen, it is critical to be confident that one can detect a mutant phenotype above the distribution of normal behaviors.
A key step is selecting a developmental age at which the behavior to be assayed is stable. Over the first week of development, several transient locomotor behaviors are manifest, including spontaneous and touch-evoked coiling [32
], olfactory aversion [17
] and cyclic swimming [34
]. Other motor behaviors mature at different time points of development. For instance zebrafish larvae have two modes of acoustic startle response distinguished by latency to movement initiation [31
]. Short latency acoustic startle responses begin to appear after 72 hpf, whereas long latency acoustic startle responses are robustly elicited after 96 hpf (Burgess and Granato, unpublished results). Similarly, a weak optokinetic response is observed at 3 days post fertilization, but becomes reliable only at 5 days post fertilization (dpf) [26
]. In an ENU mutagenesis screen, developmental delay can affect as many of 25% of families [35
]. It is therefore advisable to test fish the day after the behavior stabilizes and it is critical to exclude developmental delay as the cause of a behavioral phenotype.
Another source of developmental delay is suboptimal raising conditions. Particular care needs to be exercised over water quality in an ENU mutagenesis screen, which achieve an average of one lethal mutation per genome [35
]. One quarter of families harbor lethal mutations causing embryonic death before behavioral testing can take place. Degenerating embryos provide fodder for micro-organisms which deplete oxygen from the water. Under reduced oxygen conditions, larval development is retarded [37
]. It is therefore critical to promptly remove necrotic embryos from the pool in order to maintain water quality for the remaining fish.
However, the removal of a significant fraction of necrotic larvae from the pool introduces a source of variability into the raising conditions in that it reduces the density of the remaining fish. We found that when raised at low density, zebrafish larvae exhibit heightened startle sensitivity (A), complicating our effort to perform a 'hypersensitivity' screen. Increased startle sensitivity in low-density raised fish is specific to short latency acoustic startle responses, making it unlikely that this reflects adjustment of sensory acuity. As increased startle has been described in a variety of animals raised at low density, we were alert to the possibility of this also occurring in larval zebrafish. The multitude of factors with the potential to influence behavior underscores the importance of adhering to a strict raising protocol and noting even trivial changes to the regime.
Figure 1: Environmental manipulations alter startle responsiveness in zebrafish larvae. (A) Raising density alters startle responsiveness. Larvae were raised at a density of 3, 10 or 30 per 7 ml in a 6 cm dish, with water changes every 2 days. Startle responsiveness (more ...)
It is not always clear whether to exclude larvae with abnormal morphology from testing. Genetic pleiotropy makes it inevitable that many genes with important contributions to behavior, also act outside the brain in organogenesis and metabolism [38
]. ENU mutagenesis screens generate a high rate of visible mutations [36
] some of which, like brain necrosis or general retardation, are clearly likely to affect behavior. Nevertheless, common phenotypes like mild edema or pigmentation defects are worth including in the screen with the caveat that morphology must be excluded as the source of altered motor activity. A good example of this is the large class of mutants which fail to inflate the swim bladder [39
]. Larvae initially inflate their swim bladders by ingesting small bubbles of air. The pharynx opens by 74 hpf [40
] and later on the third day post fertilization, larvae begin to inflate their swim bladder, requiring access to the air-water interface to do so [41
]. As swim bladder inflation is itself a complex behavior requiring sensory, neural and muscular integrity, it is no surprise that a very large proportion of mutant larvae fail this task. Indeed in the Tubingen screen, 95% of mutants did not inflate their swim bladder. Similarly almost all mutants identified in a large scale insertional mutagenesis screen failed to inflate their swim bladder [42
]. Failure to inflate the swim bladder results in skeletal malformations and delayed growth [41
]. Behavioral abnormalities also result from failure to inflate the swim bladder. We found that when wildtype larvae are prevented from inflating their swim bladder by physically blocking their access to the air-water interface, they show a 50% reduction in startle sensitivity (B). Thus, it is important either to exclude swim bladder mutants from testing, or to demonstrate that failure to inflate the swim bladder does not cause a relevant behavioral phenotype.
Behavioral variability can also be introduced from the genetic background of the mutagenized stocks. While some zebrafish lines have been bred to remove embryonic lethal mutations [43
], most are not maintained as inbred stocks in order to avoid unhealthiness. As a result there is a great deal of polymorphism within zebrafish lines. Consistent with this, we find significant behavioral variability within stocks. Thus before initiating a screen, individuals to be mutagenized should be carefully tested for the behavior assayed, to ensure that the population of founders is relatively homogeneous.
Single nucleotide polymorphisms occur at a rate 10-fold higher than in other vertebrate model species [44
]. It is therefore not surprising that large variation in behavior exists between strains. For example, acoustic startle responsiveness varies significantly between our wildtype Tubingen and AB stocks (A), while prepulse inhibition of startle does not differ significantly (B). Inter-strain variability has consequences both for the design of the screen itself and for planning subsequent genetic mapping experiments [45
]. For mapping to be feasible, the wildtype behavior should be similar in at least two genetic backgrounds. It is also worth checking that the trait remains stable in the F2 offspring of the two strains, to ensure that mutation induced changes in behavior will not be masked by variability. However, even if wildtype strains show similar normal behavior, the mutant phenotype can vary on different genetic backgrounds, presumably because of differing degrees of genetic redundancy and the presence of modifier genes. We encountered this obstacle in mapping the twitch twice
mutant (Burgess and Granato, unpublished results). On a Tubingen background the phenotype was a highly penetrant set of multiple tail flips to the same side in response to a tap stimulus. On a background mixed with the WIK mapping strain, only occasional multiple same-side tail flips are seen—mutants generally respond to a tap stimulus with a ‘rolling’ startle response. Thus, strain effects on behavior do not impose an insuperable obstacle to genetic studies in zebrafish, but do demand that vigilance be exercised in designing both the screening and mapping protocols.
Figure 2: Genetic background alters startle responsiveness in zebrafish larvae. AB strain (n = 108) and Tubingen strain (n = 99) larvae were raised in identical conditions and subjected to the startle and prepulse inhibition conditions used for screening . (more ...)
Behavioral assays are notoriously sensitive to small changes in environmental conditions during testing. It is impossible to systematically test the effect of every factor that can vary in the testing protocol. Thus extreme care needs to be taken to ensure consistency of the testing arena. When renovations forced us to relocate our startle apparatus to a new room, we were aghast to find a 25% increase in startle C-bend amplitude during the first week of testing. It transpired that the difference was attributable to a 3°C drop in temperature in our new environment. Other factors requiring attention include pH, illumination and time of day. Light–dark transitions begin to entrain the larval zebrafish circadian system by the second day post fertilization with profound effects on diurnal variation in locomotor activity [46
]. It is therefore advisable to raise larvae in a light cycle incubator and pre-adapt them to a set intensity of light before beginning testing.
Before embarking on a behavioral screen, it is critical to consider what types of mutants the assay is likely to yield. Behavioral assays provide a readout of the function of multiple brain systems. For instance, we anticipated that a failure to respond to an acoustic startle stimulus could reflect abnormalities of the sensory apparatus, motor systems or central integration. It is helpful to have a battery of complimentary tests to assess the integrity of these systems in order to quickly exclude mutant lines unlikely to be informative. Even simple observations can be informative. Visually impaired fish often show darker pigmentation due to expanded melanophores [26
]. Paralysis due to disorganized muscle is easy to assess by checking muscle birefringence [25
]. Any hint of postural instability or circling behavior makes it important to examine the morphology of the inner ear and otoliths [47
]. During our screen for mutants with defects in startle modulation, we found quantification of the kinematics of movement helpful. Mutants with reduced startle amplitude also often showed reduced amplitude of tail-flips during swim bouts. This combination of features suggested that the primary defect lay at the level of the muscle or neuromuscular junction. Other mutants had defective acoustic startle responses but behaved normally to a touch stimulus, indicating that the defect was likely to represent a sensory abnormality. As we were primarily interested in modulation of normal startle responses, we excluded both these categories from further consideration.