Light Exposure And The Circadian Biology
Light exposure and circadian rhythmicity have long been known to affect the behavior and physiology of animals. When research is conducted using animal models, light intensity, duration of light exposure and the wavelength of light (usually quantified in nanometers or nm) are all considerations to be factored into experimental design. Accordingly, as with other variables, light should be handled using control and monitoring methods appropriate to the study.
As our understanding of the influence of light and circadian rhythms develop, we find that more and more sophisticated questions are being raised about the association between light, our physiology, and our health. Much of this deeper questioning has come about thanks to the expanding science of circadian biology. To help us better understand some of the mechanisms of circadian biology, Edstrom spoke with Robert Dauchy. Robert manages the Laboratory of Chrono-Neuroendocrine Oncology at Tulane University’s School of Medicine, and is also an Instructor at the Department of Structural & Cellular Biology at Tulane. He has collaborated on several published studies rooted in circadian principles, and has marshalled the effort to control and prevent light contamination in his lab and institution. We asked Robert to give us some insight regarding how light exposure can affect circadian biology, and to discuss some of the measures he and his associates have taken to control the light present in their vivarium.
THE CIRCADIAN CYCLE IN OPERATION
The 24-hour biological clock nearly always parallels the light/dark cycle of the planet. Before our Earth even had an ocean and was still-cooling molten rock, we had a light/dark cycle borne of the planet’s rotation on its axis and subsequent rotation around the sun. To think that a light/dark cycle is not important to all of the species on Earth would be an oversight.
Research associated with light/dark cycles and their biological consequences date back to the mid-1930s. Throughout the twentieth century, our awareness of the impact that light has on physiology has helped shape how we conduct experiments, and has become fundamental to how we maintain our animals’ health and well-being. Contemporary research in circadian biology is steering us toward a better understanding of cancer risk and the therapeutic responsiveness to cancer in humans, and how our habit of using light emitting devices at night may be an insidious detriment to our health.
As implied, the mechanisms involved in the light/dark cycle operate in a rhythmic loop. Here is what we understand to occur. Light can be represented as photic impulses that we receive through our eyes which provide us with our sight. Additionally, photic impulses stimulate retinal photoreceptors known as intrinsically photosensitive Retinal Ganglion Cells (ipRGCs). Unlike visual system photoreceptors (rods and cones) that are dedicated to aiding the brain with transforming light into images, the nonvisual ipRGCs serve to help regulate our circadian rhythms. The ipRGCs contain a light sensitive protein (or opsin) called melanopsin which is very sensitive to predominantly blue wavelengths of light (446-484 nm). During the daytime, the ipRGCs capture photic impulses and in turn stimulate the suprachiasmatic nucleus (SCN) of the hypothalamus. The SCN can be considered the master circadian ‘clock’. Having received this message from the ipRGCs, the SCN subsequently de-activates the pineal gland, suppressing its release of the hormone melatonin. The absence of the melatonin signal in our bloodstream is a trigger to all of our tissues that we are on a daytime cycle and amenable to daytime physiological processes. At night, when there is no light available, the pineal gland has a very robust production of melatonin which circulates throughout the body and reentrains the SCN and all of the peripheral biological clocks that its nighttime.
A BREAK IN THE CYCLE – LIGHT AT NIGHT
Light during daytime along with the nighttime melatonin signal work together to reset the suprachiasmatic nucleus - the clock which controls our day/night circadian cycle. The resulting rhythm of hormone production, as well as the production of other constituents such as arterial glucose, lactic acid and other metabolites of our bodies, functions on this pattern as entrained by the light/dark cycle. This pattern met with little interference prior to the advent of electric light.
Before electrical lighting, we were more of an agricultural society exposed to high-blue emission sunlight during the daytime and virtually no light at night or LAN. Today, this reality has changed. We are generally indoors for longer periods during the daytime and nighttime, and are regularly exposed to broad-spectrum lighting: mostly Cool White Fluorescent (CWF) and Light Emitting Diode (LED) in origin. With this consideration in mind, it’s somewhat easier to understand how normal mammalian circadian rhythms may be disrupted leading to increased incidence of human cancer and metabolic syndrome, which are now revealed in over 20 major epidemiological studies of night shift workers. Our work provides bench top laboratory animal science/biomedical research in support of these epidemiological studies.
We have been conducting studies having to do with light that is below a certain threshold for a given species. We refer to this as dim light at night, or dLAN. To a rodent that is very sensitive to light at both daytime and nighttime, exposure to dLAN as little as 0.2 lux (0.08 µW/cm2) for a minute or less is sufficient to alter a physiological response. For context, this is 20-30 times less intense than a night-light, and yet it can bring about a chain of events that disrupt natural day/night circadian regulation.
It is currently believed that exposure to LAN disrupts host/cancer circadian regulatory dynamics to include neurohormones, aerobic glycolysis (known as the Warburg effect), lipid signaling, and tumor growth prevention. Insofar as treatment of human breast and prostate cancers are concerned, it appears that LAN inhibits the circadian melatonin signal, driving intrinsic resistance to certain drug and chemotherapeutic agents. This places light exposure at night suspect of both suppressing the body’s natural means of restraining cancer, and our therapeutic means of treating it.
LAN IN THE VIVARIUM
Understanding the impact that light has on animal physiology, we realized that our research environment must be light-sanitary. So what are some of the important points to consider when deciding on lighting for an animal facility? Certainly the following are considerations: lighting efficiency (high illuminance per watt of power); lower heat production; superior spectral control; and obvious long-term lower costs. Light intensity, spectral quality, and duration of light exposure impact virtually every biological process associated with animal physiology and metabolism. Hence, appropriate lighting and lighting protocols influence animal health and wellbeing and the outcome of scientific investigations.
In our lab, all animal rooms have been completely LAN-decontaminated. Translucent observation windows were addressed to prevent light spill, and lighted in-room electrical equipment was removed. Leaky door jams, sills and frames have all likewise been modified – either with changes to the physical structure or with the application of weather stripping. In addition, exterior blackout curtains and sealed door frames were installed years ago prior to the initiation of our work.
We developed lighting protocol SOPs and monitor LAN in all animal rooms, along with daytime lighting intensities, on a daily basis. The most basic, fundamental protocol is to be attentive to the light/dark cycle, wherever you are. If you are in a dark cycle, do not walk into the room and expose the animals to light at night. Try to carry out any activity that requires animal involvement during their daylight period. If you are using red light, be aware that if it is of high enough intensity and used for a long enough exposure, it too can cause a disruption in circadian rhythms.
LIVING WITH LAN
Many people, especially those who either work during night shifts or do so periodically, can fall victim to the effects of LAN exposure. What tends to happen is that within a matter of a couple of days, established circadian rhythmicity is breached. People start to not feel well. There are overt symptoms like dysregulation of the gastrointestinal tract and interrupted sleep cycles. For individuals who are committed to night shift work, what the epidemiological studies have been showing (and what our work has been supporting) is that there are much higher rates of cancers of all types. For example, we are seeing a 60% to 80% increase in the incidence of breast cancer in night shift environments.
If someone works during a nighttime schedule, there are measures that can be taken to lessen the harmful effects of LAN. Getting a good 8 (or more) hours of sleep in a light decontaminated environment will help. Take steps to shut off lights, draw blinds, use blackout curtains, and even consider using a sleeping mask. When you are asleep, be sure that light-tight conditions are maintained. Even with your eyes closed, light of sufficient intensity can pass through the eyelids and activate the intrinsically photosensitive retinal ganglion cells of the retina, passing on photic information to the suprachiasmatic nucleus and compromising the natural cycle.
During the daytime, it’s recommended to expose oneself to more blue-enriched light, such as we receive in the normal daylight. If you are inside, try to use blue-enriched LED light as much as possible. Current research suggests that wavelengths of blue-appearing light during daytime strongly reinforce circadian rhythmicity.
Reverse this exposure at nighttime with light high in longer wavelength emission LED lighting, toward the red spectrum. Change out clocks and night-lights to red-appearing, low-intensity light. Researchers and engineers are now in the process of developing computer screens, cellular phones, tablets, TVs, and even watches which block high-blue emission light at night. The lighting industry is also developing new lights for the home, workplace, and, interestingly, in space flight to include LED lights that are high in longer wavelength LAN, or lighting that gradually changes throughout the day to mimic the outside natural light conditions at a given latitude.
As the depth and breadth of our study of circadian physiology and biorhythms flourish, many exciting possibilities are taking shape. We are making more informed choices about how we can moderate light exposure in the vivarium and in our daily lives. The effects of light and its role in circadian biology are becoming better understood, but there is still much work to be done.
Edstrom offers our sincere thanks to Robert Dauchy MS, RLATG, CMAR for providing the information discussed in this article. If you have questions regarding light monitoring in your vivarium, please contact your Edstrom Sales Representative or call us at 1-800-558-5913.
LIGHT CONTROL AND MONITORING FROM EDSTROM
For over 20 years, Edstrom has helped to support hundreds of facilities with their light control and monitoring needs. Edstrom light control and monitoring offers:
- Flexible lighting schedules for each room
- Calibrated light sensors that confirm if the lights are "Off", "On" or "On High" in the room
- Light state notification and alarming
- Built-in self diagnostics to assist with validation
Baylor College Of Medicine Transitions Mice From Bottled To Automated Watering
Automated watering systems provide facilities with many advantages over bottled watering, including reduced instances of ergonomic injury of staff, labor savings, and the reduction of long term costs. Researchers studying mouse models that use automated watering enjoy the benefit of knowing that the water the mice ingest is of a consistent, high quality. For mice that have transitioned to automated watering, there is ready access to fresh, clean water on demand.
Once the decision has been made to adopt automated watering in the vivarium, procedures need to be put in place to ensure that the transition from bottles is as smooth as possible for all involved. This includes training for animal care staff, researchers, and of course, the mice themselves. We had the pleasure of speaking with Teresa Neubauer, Operations Manager at the Baylor College of Medicine - Center for Comparative Medicine, who discussed with us how she and her team are proceeding with making the transition to automated watering.
GETTING THE MICE ACCLIMATED
Mice that are transitioning from bottles to automated water are going to need to learn a new skill, namely, finding the watering valve and drinking from it. Mice who are accustomed to drinking from a water bottle are used to finding a small droplet of water clinging to the opening at the end of the sipper tube. This clues them in as to where to drink. The automated watering drinking valve does not drip or drain on its own by design, so that telling droplet of water will not be present unless the stem of the valve is toggled. Toggling the valve is done by moving the stem of the valve laterally to the left or right. By doing this, the mice will be better able to find the water.
Just as people have differences in learning acuity and adaptiveness, so do mice. Although most mice do adapt to the new water source quickly, it will be easier for some than others. Weanlings, older mice, mice that are singularly housed and neurological mutants in particular may find the transition more challenging and will require attentive monitoring to gauge their progress. In an effort to aid the transition, all mice will have access to both a water bottle and the automated watering valve for a minimum of one week while they acclimate to their new water source.
A CLEAR MEASURE OF PROGRESS
Here at the Center for Comparative Medicine (CCM) we are using a three tiered system of color coded cards as a helpful tool to monitor and record the progress of the transition that the mice are making. Each color coded card provides our animal care staff with clear "If/Then" direction on what steps to take based on their observations during health checks. The colors that we have chosen are red, yellow and green. Here’s how they breakdown:
Red cards are the first color used and represent the beginning of the transition period for the mice. With this card, we start by recording the date that the transition period begins and the location data. During the next regular bottle change out period, we observe the bottle to see how much water is remaining in it. We do this because as time passes and the animals learn to drink from the valve, the water bottle will be utilized less and less for drinking. If the bottle is less than half full, the animals are clearly still relying on it for water and not using the valve. A new water bottle is placed on the cage and the red monitoring card is left in place.
On the other hand, if the bottle is more than halfway full, that is an indication that the valve is being put to use for drinking and the bottle can be removed from this cage. Progress! The mice have graduated to the second tier – the yellow transition card.
With the placement of a yellow transition card, we know that the bottle is out, the training wheels are off and these cages will require close attention. We record the date that the bottle was removed on the card and the location data. We also confirm that a daily hydration check is performed to ensure that the animals show no evidence of being dehydrated.
Observable, physical indications of dehydration include a hunched posture, a rough appearance to the animals’ coat and sunken eyes. If any of these conditions are present, we replace the water bottle, add moist food to the bottom of the cage, and the incident is reported. Our staff are trained and required to place an emergency call if the animal appears moribund. Staff will also observe cage bottoms for urine and feces as an indicator that the mice are drinking from the valve.
If the animals appear healthy at the time of the cage change out, we will remove the yellow transition card and replace it with a green transition card. This is our indication that the animals in this cage have successfully acclimated to automated water.
ORIENTING OUR RESEARCHERS – CLICK, TUG AND TOGGLE!
Part of our orientation for researchers involves fielding some of the particular questions they have regarding use of the automated watering system. On occasion, they may need to keep the animals on bottled water because they are involved with a study that requires specialized treatment of the water, or they may need to place the animals on water restriction. In these cases, we ask that they remove the valve from the rack and place it in a valve tray labeled ‘dirty valves’. Once the animals return to using the valve, we request that they take a valve from our ‘clean valves’ tray and add it back on to the rack stall where the original valve was removed. Our catch phrase for those who are new to placing a valve on the rack is "Click, Tug and Toggle!"
We instruct that in order to place a clean valve back on to the rack, you must first push back the manifold sleeve away from you and hold it in position. Next, slip the valve into the manifold sleeve and then release the sleeve which should make an audible click once it’s securely connected. Give the valve a gentle tug to be sure that it is properly in place. Finally, we teach that the valve should be toggled to confirm that water comes out and it is operating properly.
With a clean valve in place and functioning, we demonstrate how to determine whether the cage is properly docked in the rack. This is critical, because if the cage is not docked on the rack then the valve has very likely not passed through the cage grommet and the mice will not be able to access it in order to drink. Cage docking is aided by observing the handy ‘red dot’ indicator on the latching mechanism of the rack stall. If the red dot is visible on the latch, the cage is not docked properly. If the cage is in its proper docked position in the stall, the metal tab will cradle the plastic lug on the cage and the red dot on the latch will no longer be visible .
Acclimating both mice and personnel to an Automated Watering System can pose some challenges. As with the use of any new tool, there is a learning curve involved with transitioning from bottles to the watering valves. Taking the necessary steps of planning and training will help ease all parties through the change.
Edstrom greatly appreciates Teresa Neubauer for providing the insights presented in this article. If you have questions regarding making the transition from bottles to an Automated Watering System, please contact your Edstrom Sales Representative or call us at 1-800-558-5913.
Anatomy Of An Icon – The A160
The Edstrom A160 valve has an iconic presence in the vivarium. It is often the element that people think of when you mention automated watering for mice and rats. The A160 is purposefully designed to aid in enhancing modern Individually Ventilated Caging Systems (IVCs) in several key ways, including the elimination of repetitive stress injuries caused by daily manual bottle processing, reducing labor costs and minimizing stress to animals through less frequent cage handling. As IVCs have become a value added commodity to research facilities, automated watering systems utilizing the A160 valve have themselves presented a value added commodity to IVCs.
THE INNER WORKINGS OF A WORLD CLASS DRINKING VALVE
Throughout the years the core components of the A160 have remained true to the original, proven Edstrom design.
Guard: The guard is designed to protect the stem from unintended actuation by the animals or objects being pushed up against it. The recessed opening of the guard also helps to prevent bedding from entering the valve.
Shield: Positioned just behind the guard, the silicone rubber shield surrounds the stem, creating a flexible barrier that prevents bedding from lodging in the barrel of the valve. When an animal moves the stem, the shield moves with it and creates a small gap that allows for water to flow past it for drinking.
Stem & O-ring: A flat headed stem allows for a greater closing force as the diaphragm presses the stem head against the o-ring. This is the same leak-proof design that has been the heart of the Edstrom drinking valves for years. The eight-hole diaphragm captures the head of the stem to keep it centered.
Diaphragm: Provides the spring force to hold the stem against the o-ring. When animals release the stem, the diaphragm immediately closes the valve and water flow stops.
Cap / Check Valve: Keeps debris from entering the valve when disconnected from manifold.
MAKING THE CONNECTION
The A160 valve is able to connect to the rack watering manifold in a variety of ways. The Easy Connect version of the A160 valve is permanently mounted to the animal cage and uses an Edstrom Easy Connect Coupler to attach to the manifold piping. In A160 valves that are Affixed to the Manifold, the valve is permanently joined to the rack manifold, and requires that cages use a spring-loaded grommet door to allow the valve entry when a cage is docked on the rack. A Locking Quick Disconnect (QD) also requires the use of a grommet door on the cage for the valve to enter. In this configuration, the valve can be released from the manifold for sanitization by withdrawing the collar of the quick disconnect.
To learn more about the Edstrom A160 drinking valve and valve care, go to our website www.edstrom.com/drinkingvalvecare/ or contact an Edstrom representative.
Q: I have purchased a chlorine test kit to monitor the chlorine concentration in my automated watering system. What level of chlorine is considered acceptable for drinking water? Also, what is the best location for me to draw a sample from the system?
A: When testing chlorine levels in drinking water, a result of 2 to 3 parts per million (ppm) of free chlorine has been found to be effective at destroying upwards of 99% of bacteria, though a range of 0.5 to 10 ppm is considered acceptable.
Samples should be taken from the farthest point in the system away from the storage tank and evaluated whether concentrations are on target or need adjustment.
Q: Is there a specific type of rodent bedding material that I should use with the automated watering system, and how much of it should be added per IVC cage?
A:Bedding should be dust-free, free from irregular thin splinters or needle shaped particles, and be of a particle size greater than 0.098 inches (2.5mm). An average depth of 0.25 inch (6mm) of bedding material covering the floor of a cage is recommended.
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