Optimizing Behavioral Experiments: Key Factors Affecting Reliable Results

Optimizing Behavioral Experiments: Key Factors Affecting Reliable Results

Conducting behavioral experiments requires careful consideration of various factors to ensure reliable and reproducible results. In addition to experimental manipulations like genetic alterations and drug administration, researchers must address five critical factors influencing animal behavior studies. Understanding and managing these factors can significantly enhance the validity and reproducibility of your animal model studies. Here, we’ll explore each of these factors and provide insights on how to effectively manage them.

Manage Environment for Enhanced Habituation in Behavior Studies

The success of behavioral experiments is heavily influenced by environmental conditions. Adhering to international standards for cage size, room temperature, humidity, food, water, and bedding replacement is crucial. For example, variations in lighting environments within cage racks can impact experimental results. Researchers should carefully record and control the location of animals within the rack, especially in relation to lighting. Environmental noise, such as ventilation systems, also plays a role, requiring researchers to understand and manage differences between living and experimental areas. Ideally, however not always practical, the experimental environment should closely replicate the home cage environment.

Handle Animals Appropriately to Minimize Stress

Regular handling of laboratory animals is common practice, but behavioral investigators must take extra precaution. Habituating the animals to the investigator by handling animals daily, starting at least a week before the experiment, can mitigate unnecessary anxiety and fear and ensure more accurate data. During transitions from cages to experimental settings, gentle and prompt handling is vital. Avoiding chasing, grabbing, and minimizing midair hanging by tails is essential for maintaining the well-being and reliable behavior of the animals.

Pre-Experiment Episodes and Waiting Periods

Introducing a holding area and a specified waiting time before the experiment can reduce data variability. This strategy ensures uniformity in the episodes leading up to the experiment for all animals. The holding area, secluded from the investigator’s view, should offer a quiet environment with consistent lighting to avoid disrupting the animals. Transferring the animals from the home cage environment to the experimental environment can be stressful. Allowing for at least 30 minutes in the holding area will allow the animals to calm down following room transfer. This structured approach contributes to more consistent and reliable behavioral outcomes.

Timely Execution of Experiments

Consistency in the timing of experiments is crucial, especially for studies conducted over multiple days or involving repeated training sessions. This is particularly important as many investigators run behavioral assays during the animal’s dark (night) cycle, maintaining the same schedule each day will minimize disruptions to the animal’s sleep and increase the consistency of their performance on a given task. Following a rigid experimental schedule helps maintain stability and reduces potential confounding variables, contributing to the accuracy and reproducibility of results.

Maintaining Consistent Experimental Conditions

Acute attention to experimental conditions is also imperative. Aim to keep variables such as temperature, humidity, sound, and light consistent. Many research labs can be noisy environments, so it is important to consider the use of soundproofing solutions, such as soundproof boxes or rooms, to minimize external influences. However, be mindful of creating environments that are too silent, as this can induce unnecessary anxiety. Providing background sounds resembling the animals’ normal living environment can strike the right balance between control and ecological validity.

By addressing these five key factors, researchers can optimize the well-being of their animals and conditions of their behavioral experiments, leading to more reliable, valid, and reproducible outcomes in animal model studies.

If you would like to download these factors in a convenient application note format or are interested in learning more about additional behavioral neuroscience applications and other techniques such as Fiber Photometry and Optogenetics, please take a look at our application note and eBook page on our website.

How to Set Up an Open Field Test for Animal Behavior Research

How to Set Up an Open Field Test for Animal Behavior Research

If you are interested in studying the emotional or motor activities of animals, such as anxiety, exploration, or locomotion, you might want to consider using an open field test. This is a simple and widely used method that involves placing an animal in a large, empty box and observing its behavior. In this blog post, we will give you some tips on how to choose the right dimensions, materials, and light conditions for your open field test.

Dimensions

The size of the experimental box should be large enough to allow the animal to move freely and explore the environment, but not so large that it becomes stressful or overwhelming. The box should also be higher than the home cage to prevent the animal from escaping.

For mice, a suitable box size is at least 400 x 400 x 300 (h) mm, but you can also use a larger box, such as a 500 x 500 x 300 (h) mm box, which is ideal for open-field tests in mice. For rats, the standard box size is 900 x 900 x 500 (h) mm, but you can also use a smaller box, as long as it is larger than 600 x 600 x 500 (h) mm.

Materials

The inner walls and floor of the box should have a matte finish to reduce light reflection and glare, which can affect the animal’s vision and behavior. The box should also have a non-slippery surface to provide adequate grip for the animal and prevent falling or sliding.

While some animals may prefer wood floors and show more activity on them, wood is not a good material for open field tests because it can absorb odors and germs and cannot be easily cleaned or disinfected. Therefore, we recommend using plastic or metal boxes that can be sanitized after each use.

Another factor to consider is the transparency of the box walls. Some researchers may prefer transparent walls to observe the animal better, but this can also expose the animal to external visual stimuli, such as the experimenter, other animals, or objects, which can influence its behavior. Therefore, we recommend using opaque walls or covering the transparent walls with curtains or screens to create a more isolated and controlled environment. We also suggest using matte light gray walls and floors, as this color can help the animal acclimate faster to the new setting.

Light Conditions

The amount and quality of light in the box can also affect the animal’s behavior. Generally, animals tend to be more active and exploratory in dimmer light conditions, as they feel less exposed and threatened. In contrast, brighter light conditions can induce more anxiety and inhibition in the animal, as they feel more vulnerable and exposed.

In recent years, some researchers have reduced the light intensity in their open field tests to less than 10 lux, which is equivalent to a dark room with a faint light source. They have found that this low light level can increase the animal’s movement and exploration, compared to the traditional 50 to 200 lux, which is equivalent to a well-lit room or a cloudy day.

However, it is important to ensure that the light is evenly distributed in the box, as any contrast between light and dark areas can create a preference for the dark areas, where the animal feels more secure and hidden. To achieve uniform illumination, we recommend using LED light panels with dimming functions that can be placed on top of the box. This method can also allow you to adjust the light intensity according to your experimental needs.

Sound Environment

One of the challenges of conducting an open field test is to control the background noise in the room where you perform the experiment. Noise can interfere with the animal’s behavior and affect their responses. For example, sudden or loud noises can startle them and make them more anxious or less active.

To avoid this problem, you need to create a consistent and neutral sound environment that masks other noises in the room. One option is to use a white noise generator that produces a constant and uniform sound of 60 decibels. This can help to reduce the variability and unpredictability of the noise level and create a more stable and calm atmosphere for your animal subjects.

However, if you cannot provide the same sound environment every day, or if there are other sources of noise that you cannot eliminate, you may need to use a soundproof box or place the experimental box in a soundproof room. This can help to isolate your animal subjects from any external noise and ensure a more controlled and quiet environment for your experiment.

Experiment Duration

Another factor that you need to consider is how long you should run your open field test. The duration of the test can affect the quality and quantity of your data, as well as the welfare and comfort of your animal subjects.

Some older papers suggest that an open field test should last for 5 minutes, but this may not be enough to capture the full range of your animal’s behavior and activity. More recent studies have shown that movement often increases after the first 5 minutes, as your animal subjects become more familiar and comfortable with the environment. Therefore, it is recommended that you extend your open field test to at least 10 minutes, to allow for more exploration and adaptation.

However, if you want to measure more long-term effects or changes in your animal subjects’ behavior and activity, you may need to run your open field test for longer periods, such as 2 hours. This can help you to observe how your animal subjects respond to prolonged or repeated exposure to the same environment, and how they develop habits or preferences over time.

But remember, the longer you run your open field test, the more you need to take care of the animal’s well-being and comfort. You need to provide them with adequate food, water, and bedding, and monitor them for any signs of stress, fatigue, or injury. You also need to ensure that the temperature and humidity of the room are appropriate and consistent.

Experimental Protocol

To start the test, you need to place the animal in a specific spot in the box and let it roam freely for a certain amount of time. It is important to be consistent and use the same spot and posture for all animals, so you can compare their results fairly. A good spot to use is near the wall, because animals that are nervous or anxious tend to stay close to the edges, while those that are more confident or adventurous venture into the center.

As the animal explores the box, you can observe and record various aspects of its behavior, such as how much it moves, how often it turns, how long it stays still (freezing), and how many times it stands up on its hind legs(rearing). These indicators can tell you a lot about the animal’s emotional state, curiosity, and motivation. For example, animals that move a lot, turn frequently, and rear often are usually more active, alert, and interested in their surroundings, while animals that move little, turn rarely, and rear less often are usually more passive, calm, and cautious which may indicate an anxiety-like state.

To increase the accuracy and convenience of data analysis, you can use a more advanced Open Field system, which automatically measures and analyzes the animal’s behavior using image processing and/or infrared beams. This approach also allows for automatic detection of rearing. However, keep in mind that different researchers may have different interpretations of what these behaviors mean, so you should always focus on the changes in the frequency and duration of these behaviors, rather than their absolute values.

After the test is over, you need to remove the animal from the box as quickly as possible, to avoid stressing it or affecting its behavior. You also need to clean and disinfect the box before testing the next animal. This is to prevent any contamination or odor that could influence the animal’s behavior. If you are using mice, especially the C57/B6 strain, you should avoid using alcohol to disinfect the box, because they are very sensitive to it. You should wait until the alcohol evaporates completely, or use a different disinfectant, before testing the next mouse.

Summary

While the Open Field test method remains an invaluable tool for researchers studying a wide range of behaviors. It is important to take the proper measures to reduce environmental and experimenter bias in your results. By following the above guidelines, you can conduct open field tests with precision and reproducibility, ensuring the collection of reliable data on animal motor function and emotional states. If you are interested in learning more about additional behavioral neuroscience applications and other techniques such as Fiber Photometry which can be paired with behavioral assays, please take a look at our application note and eBook page on our website.

​​Fluid Intake Precision: The Drinko Advantage

​​Fluid Intake Precision: The Drinko Advantage

Introduction

Precise fluid intake measurement can be extremely important in behavioral neuroscience studies. Accurately measuring the amount of fluid a rodent consumes provides crucial insights into their physiological and behavioral responses. In this blog, we will delve into the significance of precise fluid intake measurement, highlighting the Drinko Measurer. With its enhanced features like precise and accurate drink measurement, leak-free sipper tube, and multiple sizes designed to fit, the Drinko Measurer has improved the way researchers track and analyze fluid consumption in rodent studies.

The Importance of Accurate Fluid Intake Measurement

Accurate measurement of fluid intake is vital in behavioral neuroscience studies for several reasons. Firstly, it allows researchers to evaluate the impact of specific treatments, drugs, or interventions on an animal’s drinking behavior. By precisely quantifying fluid intake, researchers can analyze the effects of these factors on hydration levels, metabolism, and overall physiological state. Secondly, precise measurement enables the detection of subtle changes in drinking behavior that may be indicative of altered cognitive or emotional states. This information is crucial for understanding underlying behavior and for drawing accurate conclusions in neuroscience research.

The Drinko Measurer: Leak-Free Fluid Intake Measurement

The Drinko Measurer has emerged as an ideal tool in the field of behavioral neuroscience for accurately measuring rodent fluid intake. This innovative device provides numerous benefits that enhance the accuracy of fluid intake measurement. Equipped with a double-ball bearing, the Drinko Measurer offers precise volume measurement, ensuring reliable data collection every time. Researchers can track and record the exact amount of fluid consumed by rodents, enabling precise analysis and comparison between experimental groups.

Explore the Drinko Measurer for Precise Fluid Intake Measurements!

The Advantages of the Leak-Free Sipper Tube

The Drinko Measurer incorporates a leak-free sipper tube system, addressing a common challenge in fluid intake measurement. With conventional setups, fluid leakage can lead to inaccuracies and potential confounding factors. However, the Drinko Measurer’s leak-free sipper tube eliminates this issue, ensuring that the measured fluid intake truly represents the consumed amount. This feature enhances the reliability of the data and allows researchers to draw more accurate conclusions regarding the effects of various treatments or experimental conditions on fluid intake.

Versatility and Adaptability with Multiple Sizes

The Drinko Measurer offers researchers the flexibility to adapt to different experimental requirements by providing multiple sizes of sipper tubes. Whether researchers are working with mice, rats, or other rodent models, they can select the appropriate sipper tube size to ensure accurate measurement. This adaptability makes the Drinko Measurer a valuable tool for various behavioral neuroscience studies.

 

Conclusions

Accurate measurement of fluid intake is essential in behavioral neuroscience studies, as it provides valuable insights into the physiological and behavioral responses of rodents. The Drinko Measurer, with its accurate drink measurement, leak-free sipper tube, and multiple sizes, has revolutionized fluid intake measurement. By utilizing this advanced device, researchers can ensure precise data collection, eliminate confounding factors, and enhance the reliability of their findings in behavioral neuroscience research.

Touch Panel Operant System for Neurological Deficit Research

Touch Panel Operant System for Neurological Deficit Research

1: Introduction and Background

Both visual discrimination (VD) and reversal learning (RL) in neurological studies are very important to study animals’ cognitive abilities.   

  • Reversal Learning:  

Reversal learning requires an animal to learn to discriminate two different stimuli but reverse its responses to these stimuli every time it has reached a learning criterion. Thus, different from pure discrimination experiments, reversal learning experiments require the animal to respond to stimuli flexibly, and the reversal learning performance can be taken as an illustration of the animal’s cognitive abilities. (1)

  • Visual Discrimination:

As for visual discrimination task, which is a task used extensively in the elucidation of cognitive impairment produced by lead, is a test in which the formerly correct stimulus becomes the incorrect one, and vice versa. (2)

2: Advantages of Touch Panel Operant System in Reversal Learning and Visual Discrimination

Our Touch panel operant training system has standard mazes that come with video tracking and automated data collection and analysis. It also contains several key features that make it a wise choice for rodent behavior, specifically mouse behavioral testing. The unique trapezoidal design creates a space for animals to focus more on the screen ahead during rodent behavior testing.

 

The Touch Panel operant training system contains infrared (IR) sensor technology, which locates at the top of the touchscreen, to improve the accuracy with which the system can detect touch responses from mice and rats. Unlike the touch screen technology that most of our smartphones and computer screens use, called projective capacitance, the infrared sensors inside the touch screen improve accuracy by eliminating the need for a minimum force required to generate a response. This means that even nose poke’s from mice will register a response.

TouchPanelScreen
Touch Panel Operant Chamber

In addition to improved sensitivity, our chamber was designed in a trapezoidal shape instead of a square, making it easier for the animal to focus on the screen ahead. Our touch panel chambers are compatible with in vivo electrophysiology and optogenetics techniques, as well as with miniature head-mounted microscopes. 

The system also includes software that enables users to design and run their own tasks with video tracking capabilities for automated data collection.The chambers come with our Operant TaskStudio Software package, an extremely user-friendly software platform that enables customers to design and execute their own tasks or choose from a variety of pre-programmed tasks.

operant-taskstudio

3: Example of Research

The team of Tatsuhiro Ayabe, Rena Ohya and Yasuhisa Ano used our touch panel-based operant system integrated with visual discrimination (VD) and reversal learning (RD) protocols.

During the VD task, vertical and horizontal stripes were shown on the screen as visual stimuli where vertical stripes were used as the correct response to half of the mice while horizontal stripes were used as the incorrect response to the other half of the mice (Figure 1). A trial started when the mouse touched the reward magazine. If the mouse was recorded poking at the vertical stripes (correct stimulus), it would be rewarded, and a 2-second inter-trial interval (ITI) would follow up. If the mouse poked at the horizontal stripes (incorrect stimulus), there would be no reward and a 5-second darkness would take place, followed up by a 5-second ITI. If the mouse touched the reward magazine again after each ITI, a new trial began. If the mouse failed to poke either stimulus in 30 seconds, the trial would be pruned. 60 minutes before the test session, the Iso-α-acids solution (1 mg/kg body weight) or distilled water (DW) would be administered through oval gavage. 30 minutes before the test session, scopolamine (0.8 mg/kg body weight) or saline would be intraperitoneally administered (Figure 2). Mice were expected to have a correct response rate that was above 80% (post-VD training) so that they would perform the VD test without drug treatments until that correct rate was achieved before they started performing the RD task.

                                Figure 1

Figure 2

After the treatment of scopolamine and iso-α-acid, both treated groups would perform the RD task. For the RD task, the visual stimuli were switched in the opposite order compared to the VD task, where horizontal stripes were used as the correct response and vertical stripes were used as the incorrect response. Only Iso-α-acids solution (1 mg/kg body weight) or distilled water (DW) would be administered through oval gavage 60 minutes before the test session for the RD task, and the scopolamine would not be administered to mice. DW and IAA solution was administered 17 times and scopolamine was treated 7 times in total during all experiment periods. The number of correct response changes was obtained by calculating the difference between the correct rate of each daily trial and the correct response rate of the first trial so that the efficiency of mice changing their previous memory conditions could be evaluated.

See the full publication

4. Applicable tasks for the Touch Panel Operant System

Task:Description:Measures:Useful for studying:
Visual Discrimination (VD)Subject learns that one of two shapes is the correct stimulus that results in food/liquid reward.Correct response is then changed-reversal learning (RD).

Cognitive flexibility

 

 

Neuropsychiatric disorders (Schizophrenia, Autism)

Research Example 1: β-lactolin, a whey-derived glycine―threonine―tryptophan―tyrosine lactotetrapeptide, improves prefrontal cortex-associated reversal learning in mice

Research Example 2: Hop-Derived Iso-α-Acids in Beer Improve Visual Discrimination and Reversal Learning in Mice as Assessed by a Touch Panel Operant System

Paired Associate Learning (PAL)Subject must learn and remember which of 3 objects goes in which correct spatial location. Each trial involves 2 objects, 1 in the correct place and the other incorrect. Mice must choose the correct object for reward.Hippocampal dysfunctionNeurodegenerative diseases (Alzheimers and Dementia)
Visuomotor Conditional Learning (VCL)Stimulus-response task. Subject must learn that two stimuli go with two different locations. When a stimulus is presented, the subject must respond to the location associated with specific stimulus.Motor dysfunctionMotor disorders (Parkinsons and Huntingtons disease)
5-Choice Serial Reaction Time (5CSRT)Subjects must respond to brief visual stimuli presented in 1 of 5 locations. Stimulus disappears after a set interval which requires the subject to return the location by memory.Attention span and impulsivityAnimal models of ADHD and Schizophrenia

Explore Our Touch Panel Operant System!

References:

(1): Bublitz, Alexander, et al. “Reversal of a Spatial Discrimination Task in the Common Octopus (Octopus Vulgaris).” Frontiers in Behavioral Neuroscience, vol. 15, 2021, https://doi.org/10.3389/fnbeh.2021.614523.

(2): Slikker, William, and Cheng Wang. “Chapter 31.” Handbook of Developmental Neurotoxicology, Academic Press, 1998, pp. 539–557. https://doi.org/10.1016/B978-0-12-648860-9.X5000-6

(3): Ayabe, T., Ohya, R., & Ano, Y. (2019). Hop-Derived Iso-α-Acids in Beer Improve Visual Discrimination and Reversal Learning in Mice as Assessed by a Touch Panel Operant System. Frontiers in Behavioral Neuroscience13, 67.

Use head-fixation to identify neurons associated with voluntary movements

Use head-fixation to identify neurons associated with voluntary movements

1. Head-Fixation in Neuroscience Research

Head-fixation is widely used in the field of behavioral neuroscience in awake behaving animals to better understand the cortical involvement in animal behaviors. Our understanding of topics such as associative learning, sensory perception, navigation and motor control have been greatly improved by head-fixed experimental preparations. The restraint and motion minimization of the animal’s head minimizes noise and motion artifacts often seen in freely moving studies, which allows for stimulus control studies, perturbation experiments, neural recordings and in vivo cellular imaging. It also simplifies the experimental set-up design and data analysis, allowing for chronic and long-duration studies as many systems facilitate accurate repeated alignment between the animal’s head and the restraining system.

Most recently, head-fixation has been used alongside advanced techniques such as high-density electrophysiology recordings and two-photon imaging to investigate neural circuits in vivo that would not be possible to study in freely moving subjects.

2. Limitations to Most Head-Fixation Systems

However, there are a number of limitations that need to be considered when carrying out large-scale monitoring of neuronal activity under head-fixed operant conditions.

  • Many models of head-fixation assume that head movements don’t occur when the animal is restrained. Some subtle movements of the animal’s head that cannot be easily observed or taken into consideration when doing data analysis can result in noise or artifact production. This can lead to significant confounding variables, with a misinterpretation of neural activity associated with animal behavior. Choosing the correct head-fixation instrumentation reduces the risk of confounding variables associated with involuntary head movements.
  • Measuring motor responses in rats or mice by monitoring whisker movements and licking behavior have been used in conditional behavioral tasks in the past. However, these motor movements are driven by a pathway in the brain involving automatic repetition that involves the brainstem. Identifying a reliable, measurable movement under operant learning conditions is imperative to accurately associate cortical activity with behavior.

3. Skilled Motor Movements in Operant Conditioning

Skilled motor movements are a good measure of operant learning, as rats and mice are naturally able to perform voluntary skilled motor movements using their forelimbs. Skilled movements are intentional isolated movements with specific parts of the body, which require the recruitment of different neural pathways that contain different subtypes of neurons firing with synchronicity. A number of operant learning paradigms can be studied by measuring forelimb movement response under head-restraint conditions alongside high-density electrophysiology recordings or two-photon imaging. It is important to consider the type of restraining system to use when studying voluntary forelimb movement in rodents.

4. Case Study – Microcircuitry coordination of cortical motor information in self-initiation of voluntary movements

Using the TaskForcer (O’Hara) restraining system, The Isomura* group at the University of Tokyo uncovered functional diversity of pyramidal cells and the uniformity activation of fast-­spiking interneurons across all cortical layers in the expression of trained rodent voluntary movement. Furthermore, they identified a pattern of excitatory synaptic interactions among neighboring neurons that play different roles in self-initiated voluntary forelimb movement.

Experimental Procedure

Juxtacellular and multi-unit recordings were taken from the motor cortex of 74 rats who were head-restrained and trained to repeat voluntary forelimbs movements. The juxta-­cellular recording technique was implemented as it provides accurate spike events and morphological features for a cortical or subcortical neuron. The multiunit recording technique is useful for exploring the synaptic connectivity of many neurons simultaneously while remaining blind and unbiased.

Amuza TaskForcer

A multi-­rat task-­training system was developed to simultaneously train up to six adult rats on an operant voluntary forelimb-­movement task. During the operant trials, the trainee rats quickly learned the relationship between the lever and the water reward. This led to the trainee rats casually grabbing the lever with their right forelimb for their reward instead of struggling during the restraining process. 

TaskForcer Lever

All 74 rats were successfully able to perform the operant motor task in just 8 days of training. Post training, the rats were transferred to a recording room where they were able to perform the same casual operant motor task during juxtacellular and multiunit recordings. Several distinct  patterns of neuronal firing at an electrode site in relation to the forelimb-­movement task were revealed during these multi-unit recordings.

Focus on results!

When research animals are stressed or distracted, training can slow and your results are what will suffer variability. Unlike other systems that allow rodents to essentially “run free” while attempting to perform tasks at the same time, Amuza’s TaskForcer eliminates those distractions—and their effects on your data.

Resources

*Isomura Y, Harukuni R, Takekawa T, Aizawa H, Fukai T. Microcircuitry coordination of cortical motor information in self-initiation of voluntary movements. Nat Neurosci. 2009 Dec;12(12):1586-93. doi: 10.1038/nn.2431. Epub 2009 Nov 8. PMID: 19898469.

Accurately monitor reward-oriented licking

Accurately monitor reward-oriented licking

Identifying efficient robust methods to monitor licking behavior in rodents is key to
understanding the role of reward-oriented dopaminergic neural pathways in animal behavior. By
monitoring licking behavior, researchers can better understand how rodents gauge the outcome
of a specific reward, their incentive for the reward and how they predict the reward (1).

However, licking microcircuitries in the brain are complex, and incorporate a number of different
neurons controlling different behaviors. A difficult task in recent times has been accurately
identifying the specific microcircuit associated with each specific reward-oriented lick behavior.

The mesolimbic dopamine system is involved in reward-oriented behaviours, and dopamine
antagonism in rodents has been shown to change ingestive behaviors. Pharmacology DAergic
stimulation of the NAc triggers an intense response to obtain a reward, even if a rat has
undergone extinction training (1). While ingestive behaviors include both feeding and drinking,
the exact involvement in water drinking remains unclear.

As mice respond to sensory stimuli by licking for liquid rewards, precise monitoring of licking
during these tasks provides an accessible metric of sensory-motor processing, particularly when
combined with simultaneous neural recordings or microdialysis (2). The precise timing of reward
consumption is critical to understand associations between neural activity and animal behavior.
Therefore using the right detection method as well as a reliable rodent model of dopamine
ensures that licks are monitored reliably (3).

Licking Units – Limitations to be considered

Some of the main challenges when developing and implementing lick detectors during head-restraint
microdialysis or neurophysiological experiments in mice include:

    1.  Electrical contact sensors that trigger food or water feeder to dispense can create
      electrical artifacts that are similar to neural or behavioral amplitudes and time courses,
      which can also interfere with electrophysiology recordings (4).
    2. Temporal characteristics of licking (approx 7Hz) are different from the profile of individual
      licks which are much faster. This is important if trying to determine the onset/offset of
      licks. If lick is being used to send TTL signals to other devices it can be troublesome.
    3. Mice are small, so behavior can be disturbed by equipment that is bulky and obstructs
      animal view.
    4. Head-restraining animals without proper habituation increases cortisol levels and could affect neural recordings / microdialysis. There it is important to ensure adequate training time

Contactless photo-sensors such as infrared detectors overcome these obstacles when
monitoring lick behavior and remove any electrical artifact interference from the set-up.

Fig. 1: Transgenic construction of DSI mice.

Different promoters expressed in each line of transgenic model. Tamoxifen administration used to induce activation of transgenic phenotype.
Dopaminergic synaptic vesicles are prevented from releasing neurotransmitters by v-SNARE cleaving in DSI models.

Case Study
Drinking behavior was analyzed in triple transgenic mice generated with reduced DA release and treated with a D1-like or D2-like DA receptor agonist. Triple transgenic mice were generated to secrete reduced dopamine levels in the striatum and nucleus accumbent compared to control. These triple transgenic mice made fewer licks and fewer lick bursts than control under thirsty conditions. D1 or D2/3 receptor agonists were then administered to identify the influence of dopamine receptors in altered drinking behavior.

New triple transgenic mouse line expected to exhibit partial blockade of synaptic release rather than severely impaired DA secretion seen in other dopamine-depleted mice models. The DSI mouse line enables the study of phenotypes related to DA loss and the role of DAergic neurons and the DA receptors in drinking behavior.

Fig 2: Training of mice to lick for a water reward.

(a) Scheme of the training for licking test. (b) After 2 days of water deprivation, control and DSI mice were trained to lick a water nozzle for a water reward (4 μl/lick) (RM‐ANOVA:genotype, p < .05; time, p < .01). The daily water intake was limited to 1.5 ml per day, and the body weight was maintained at the same level (Ctrl, n = 16; DSI, n = 16). *p < .05 compared to Ctrl mice. Values are shown as the means ± SEMs

Experimental set-up
The apparatus for licking training and data recording includes a water-pumping device and an infrared beam detector system which are controlled by software.

Thirsty mice showed vigorous activity when water was available, and they drank from different angles either in front of or under the water nozzle. This tendency reduced the accuracy of recording. Thus, the researchers utilized an apparatus (TaskForcer, O’Hara) that monitors neural circuits while a mouse is licking. A custom-made head plate was fixed onto a mouse’s skull with dental acrylic to reduce its head movements.

Assessed drinking behavior by analyzing licking microstructure

  • Number of licks and bursts
  • Size of bursts
  • Intraburst lick speed

A burst was defined as continuous licking (>2 licks with <0.4 s between licks).

After 2 days of water deprivation, the mouse was placed inside an acrylic tube and trained to lick for a water reward for 15 mins per day for 7 consecutive days.

Each interruption of the infrared beam counted as one lick, and the mouse was rewarded with one unit of water (4uL of water per lick).

Microdialysis was carried out using equipment supplied by Amuza Inc. to monitor levels of dopamine in the brain of both control and transgenic mice.

Fig 3: Scheme of the rat licking microstructure.

(a) The number of total licks carried out represents the extent of water drinking activity and therefore reflects the general drinking behavior. The number of bursts indicates the activation of responses and thus represents the incentive motivation triggered by reward cues. (b) DSI mice made fewer licks and bursts than the control littermates. The D1 receptor agonist ameliorated the lick number but did not increase the burst number, and the D2 receptor agonist suppressed all the measurement results from the licking test. The D1 agonist A68930 was effective only for DSI mice, but the D1 agonist SKF38393 was effective for both control and DSI mice

Results
Findings suggest that D1 receptor activity impacts drinking and may also contribute to treatment
for illnesses related to DA loss.

DSI mice avoid the infirmity and reduced food and water consumption exhibited by DA-deficient
mice.

DSI mice showed impaired motor control when given a challenging rotarod test and made fewer
licks and bursts than control mice.

One D1 receptor agonist increased the number of licks made by thirsty DSI mice.
While another increased the number of licks made by DSI and control mice.

Combine Operant Tasks and Rewards

TaskForcer is the must-have modular system for in-vivo electrophysiology and imaging. Our operant-behavior conditioning system is designed around your priorities. The only animal training system that was created to speed training, simplify configuration, and deliver consistent results to expedite your discoveries.

Resources

(1) Kao K‐C, Hisatsune T.: Differential effects of dopamine D1‐like and D2‐like receptor
agonists on water drinking behaviour under thirsty conditions in mice with reduced dopamine
secretion. Eur J Neurosci. 2019;00:1–14. Ht
References:
(2) Williams, B., Speed, A., Haider, B: A novel device for real-time measurement and
manipulation of licking behavior in head-fixed mice (2018)
(3) Clark, Nicholas McKinley.: FAST, ACCURATE, AND LOW-COST SENSING OF REWARD
CONSUMPTION AND LICK RESPONSES IN RODENTS (2018)
(4) Hayar, A., Bryant, J.L., Boughter, J.D., Heck D.H.: A low-cost solution to measure mouse
licking in an electrophysiological setup with a standard analog-to-digital converter (2008)